U.S. patent number 10,758,596 [Application Number 16/546,424] was granted by the patent office on 2020-09-01 for compositions and methods to prevent and treat biofilms.
This patent grant is currently assigned to ZIOLASE, LLC. The grantee listed for this patent is ZIOLASE, LLC. Invention is credited to Brad W. Arenz, Thomas K. Connellan, Dennis W. Davis, Svetlana A. Ivanova.
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United States Patent |
10,758,596 |
Ivanova , et al. |
September 1, 2020 |
Compositions and methods to prevent and treat biofilms
Abstract
Compositions and methods to treat biofilms are disclosed based
on the discovery of the role of the disaccharide trehalose in
microbial biofilm development. In various embodiments to treat
body-borne biofilms systemically and locally, the method includes
administering trehalase, the enzyme which degrades trehalose, in
combination with other saccharidases for an exposition time
sufficient to adequately degrade the biofilm gel matrix at the site
of the biofilm. The method also includes administering a
combination of other enzymes such as proteolytic, fibrinolytic, and
lipolytic enzymes to break down proteins and lipids present in the
biofilm, and administering antimicrobials for the specific type(s)
of infectious pathogen(s) underlying the biofilm. Additionally,
methods are disclosed to address degradation of biofilms on medical
device surfaces and biofilms present in industrial settings.
Inventors: |
Ivanova; Svetlana A. (Winter
Springs, FL), Davis; Dennis W. (Palm Bay, FL), Arenz;
Brad W. (Orlando, FL), Connellan; Thomas K.
(Charlottesville, VA) |
Applicant: |
Name |
City |
State |
Country |
Type |
ZIOLASE, LLC |
Winter Springs |
FL |
US |
|
|
Assignee: |
ZIOLASE, LLC (Winter Springs,
FL)
|
Family
ID: |
60157180 |
Appl.
No.: |
16/546,424 |
Filed: |
August 21, 2019 |
Prior Publication Data
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Document
Identifier |
Publication Date |
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US 20200000889 A1 |
Jan 2, 2020 |
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Related U.S. Patent Documents
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Application
Number |
Filing Date |
Patent Number |
Issue Date |
|
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15632618 |
Jun 26, 2017 |
10420822 |
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13481787 |
May 26, 2012 |
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61520654 |
Jun 13, 2011 |
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Current U.S.
Class: |
1/1 |
Current CPC
Class: |
C12Y
302/01028 (20130101); A61K 38/14 (20130101); A61K
9/006 (20130101); A61K 31/7036 (20130101); A01N
63/00 (20130101); A61K 31/546 (20130101); C12N
9/2405 (20130101); A61K 31/496 (20130101); A61K
8/66 (20130101); A61K 38/47 (20130101); A61P
31/04 (20180101); A01N 63/10 (20200101); A61K
9/7007 (20130101); A61Q 11/00 (20130101); A01N
63/10 (20200101); A01N 25/34 (20130101); A01N
63/10 (20200101); A01N 63/10 (20200101); A61K
31/7036 (20130101); A61K 2300/00 (20130101); A61K
31/546 (20130101); A61K 2300/00 (20130101); A61K
38/14 (20130101); A61K 2300/00 (20130101); A61K
31/496 (20130101); A61K 2300/00 (20130101); A01N
63/10 (20200101); A01N 25/34 (20130101); A01N
63/10 (20200101); A01N 63/10 (20200101); Y10T
29/49826 (20150115) |
Current International
Class: |
A61K
38/47 (20060101); A61K 31/496 (20060101); A61K
8/66 (20060101); A61P 31/04 (20060101); C12N
9/24 (20060101); A61Q 11/00 (20060101); A61K
9/70 (20060101); A61K 9/00 (20060101); A01N
63/00 (20200101); A61K 31/546 (20060101); A01N
63/10 (20200101); A61K 38/14 (20060101); A61K
31/7036 (20060101) |
References Cited
[Referenced By]
U.S. Patent Documents
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1068871 |
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EP |
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JP |
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WO |
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2008004128 |
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Jan 2008 |
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WO |
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2008141416 |
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Nov 2008 |
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WO |
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Primary Examiner: Claytor; Renee
Assistant Examiner: Clarke; Trent R
Attorney, Agent or Firm: Allen, Dyer, Doppelt + Gilchrist,
P.A.
Parent Case Text
RELATED APPLICATIONS
This is continuation application of Ser. No. 15/632,618 filed Jun.
26, 2017, which is a continuation-in-part application of Ser. No.
13/481,787 filed May 26, 2012, which is based on provisional
application Ser. No. 61/520,654 filed Jun. 13, 2011, the
disclosures which are hereby incorporated by reference in their
entirety.
Claims
That which is claimed is:
1. A method of producing a tissue dressing for application to a
skin wound for reducing growth of biofilm mass and biofilm cells in
the skin wound resulting from Staphylococcus aureus or Pseudomonas
aeruginosa bacteria, comprising: providing a sterile tissue
dressing having no biofilm growth resulting from Staphylococcus
aureus or Pseudomonas aeruginosa bacteria; applying silver and
trehalase onto the tissue dressing, wherein the silver is from
about 0.1 mg/cm.sup.2 of tissue dressing surface area to about 1.0
mg/cm.sup.2 of tissue dressing surface area, and the trehalase is
from about 0.5 mg/cm.sup.2 of tissue dressing surface area to about
10 mg/cm.sup.2 of tissue dressing surface area, wherein the
trehalase is derived from a mammalian or plant source.
2. The method of claim 1, wherein the tissue dressing comprises
cotton fibers.
3. The method of claim 1, wherein the tissue dressing includes an
antibiotic selected from the group consisting of Ceftazidime,
Gentamicin, Tobramycin, Vancomycin, and Ciprofloxacin.
4. The method of claim 3, wherein the antibiotic comprises
Gentamicin.
5. The method of claim 1, wherein the silver comprises silver
particles of less than 20 nanometer diameter.
6. The method of claim 1, wherein the trehalase comprises 0.5 to
10.0 mg per milliliter on a protein basis.
7. The method of claim 1, wherein the silver and trehalase are
applied onto the tissue dressing as a gel.
8. The method of claim 1, wherein the silver comprises a sustained
release silver.
9. The method of claim 8, wherein the sustained release silver
comprises silver nitrate that does not exceed a concentration of 1%
in contact with living tissue.
10. The method of claim 8, wherein the sustained release silver
comprises silver sulphadiazine at about 1% in a cream base or
bandage.
11. The method of claim 8, wherein the sustained release silver
comprises 1% to 5% silver sulphadiazine and the trehalase is about
0.25% to about 2.0%.
Description
FIELD OF THE INVENTION
The present disclosure is generally related to compositions and
methods to prevent and treat biofilms.
DESCRIPTION OF RELATED ART
Over the last century, bacterial biofilms have been described as a
ubiquitous form of microbial life in various ecosystems which can
occur at solid-liquid, solid-air, liquid-liquid, and liquid-air
interfaces. The general theory of biofilm predominance was defined
and published in 1978 (Costerton J W, Geesey G G, and Cheng G K,
"How bacteria stick," Sci. Am., 1978; 238: 86-95.). The basic data
for this theory initially came mostly from natural aquatic
ecosystems showing that more than 99.9% of the bacteria grow in
biofilms on a variety of surfaces, causing serious problems in
industrial water systems as well as in various pipelines and
vessels.
Later this fundamental theory of bacterial biofilm was accepted in
the medical and dental areas. New and advanced methods for the
direct examination of various biofilms showed that microorganisms
that cause many medical device-related and other chronic infections
in the human body actually grow in biofilms in or on these devices,
as well as on mucosal linings of various organs and systems (oral
cavity, respiratory tract, eyes, ears, GI tract, and urinary
tract). As stated in this theory, "bacteria have certain basic
survival strategies that they employ wherever they are" (Donlan R M
and Costerton J W, "Biofilms: Survival Mechanisms of Clinically
Relevant Microorganisms," Clinical Microbiology Reviews, April
2002: 167-193.)
The Nature and Structure of Biofilms
Over decades, direct physical and chemical studies of various
biofilms (mostly grown in laboratory settings) show that they
consist of single microbial cells and microcolonies, all embedded
in a highly hydrated exopolymer matrix comprising biopolymers of
microbial origin, such as polysaccharides (the major component),
proteins, glycoproteins, nucleic acids, lipids, phospholipids, and
humic substances; ramifying water channels bisect the whole
structure, carrying bulk fluid into the biofilm by convective flow,
providing transport of nutrients and waste products, and
contributing to a pH gradient within the biofilm (Costerton J W and
Irvin R T, "The Bacteria Glycocalyx in Nature and Disease," Ann.
Rev. Microbiol., 1981; 35: 299-324.); (de Beer D, Stoodley P, and
Lewandowski Z, "Liquid flow in heterogeneous biofilms," Biotechnol.
Bioeng, 1994; 44: 636-641.); (Himmelsbach D S and Akin D E,
"Near-Infrared Fourier-Transform Raman Spectroscopy of Flax (Linum
usitatissimum L.) Stems," J Agric Food Chem, 1998; 46: 991-998.);
(Maquelin K, Kirschner C, Choo-Smith L P, van den Braak N, Endtz H
P, Naumann D, and Puppels G J, "Identification of medically
relevant microorganisms by vibrational spectroscopy," J Microbiol
Methods, 2002; 51: 255-271.); (Neu T R and Marshall K C, "Bacterial
Polymers: Physicochemical Aspects of Their Interactions at
Interfaces," J Biomater Appl, 1990; 5: 107-133.); (Neugebauer U,
Schmid U, Baumann K, Ziebuhr W, Kozitskaya S, Deckert V, Schmitt M,
Popp J, "Toward a Detailed Understanding of Bacterial
Metabolism--Spectroscopic Characterization of Staphylococcus
Epidermidis," ChemPhysChem, 2007; 8: 124-137.); (Weldon M K,
Zhelyaskov V R, Morris M D, "Surface-enhanced Raman spectroscopy of
lipids on silver microprobes," Appl Spectrosc, 1998; 52: 265-269.).
Depending on the biofilm type and the microorganisms involved,
microcolonies of microbial cells make up approximately 10%-15% of
the biofilm by volume, and the biofilm matrix comprises
approximately 85%-90%. Water, the major component of the biofilm
matrix, can make up to 95%-98% of the matrix volume, and the
particulate fraction of the matrix can comprise the rest 2%-5%
correspondingly. Extracellular polysaccharides and proteins have
been considered to be the key components of the biofilm matrix and
have been most extensively studied over decades (Sutherland I W,
"The biofilm matrix--an immobilized but dynamic microbial
environment," Trends Microbiol, 2001; 9: 222-227.); (Stewart P S
and Costerton J W, "Antibiotic resistance of bacteria in biofilms,"
Lancet, 2001; 358: 135-138.); (Staudt C, Horn H, Hempel D C, Neu T
R, "Volumetric measurements of bacteria and EPS-glycoconjugates in
biofilms," Biotechnol Bioeng, 2004; 88: 585-592.); (Zhang X Q,
Bishop P L, and Kupferle M J, "Measurement of polysaccharides and
proteins in biofilm extracellular polymers," Water Sci Technol,
1998; 37: 345-348.).
Polysaccharides, postulated to be the key component of the biofilm
matrix, provide diverse structural variations of the glycocalux
formed by saprophytic and pathogenic microorganisms in a variety of
environments (Barbara Vu, et al., "Review. Bacterial extracellular
polysaccharides involved in biofilm formation," Molecules, 2009;
14: 2535-2554; doi: 3390/molecules 14072535.). The types of
polysaccharides in microbial biofilms are of enormous range and
depend on the genetic profile of microorganisms involved and the
physicochemical properties of local environment (Sutherland I W,
"The biofilm matrix--an immobilized but dynamic microbial
environment," Trends Microbiol., 2001; 9: 222-227.). Many
polysaccharides are constitutively produced by various bacteria as
structural elements of the bacterial cell wall and virulence
factors; they can stay attached to the bacterial cell wall surface,
forming a complex network surrounding the cell with electrostatic
and hydrogen bonds involved, or they can be released into media as
exopolysaccharides (EPS) (Mayer C, Moritz R., Kirschner C., Borchar
W, Maibaum R, Wingender J, and Flemming H C, "The role of
intermolecular interactions: studies on model systems for bacterial
biofilms," Int J Biol Macromol, 1999; 26: 3-16.). Polysaccharides,
as well as mono- and disaccharides, can be taken by bacteria from
the environment and metabolized as a carbon source, and their
metabolism is genetically regulated via balanced production of
enzymes for both synthesis and degradation pathways (Sutherland I
W, "Polysaccharases for microbial polysaccharides," Carbohydr
Polym, 1999; 38: 319-328.). Depending on their structure, EPS can
bind various amount of water, and some of them (such as cellulose,
mutan or curdlan) can even exclude most water from their tertiary
structure. Over the years, the gel-like viscosity of the biofilm
matrix was attributed mainly to the physical and chemical
properties of the polysaccharides involved (Christensen B E, "The
role of extracellular polysaccharides in biofilms," J Biotechnol,
1989; 10: 181-202.); (Stoodley P, et al., "Oscillation
characteristics of biofilm streamers in turbulent flowing water as
related to drag and pressure drop," Biotechnol Bioeng, 1998; 57:
536-544.). Exopolysaccharides can be neutral homopolymers (such as
cellulose, dextrans, levans), but the majority are polyanionic (for
example, alginates, gellan, xanthan produced by Gram-negative
bacteria) with attraction of divalent cations (Ca, Mg) to increase
binding force, and a few are polycationic, such as those produced
by some Gram-positive bacteria (Sutherland I W, "Biotechnology of
Exopolysaccharides," Cambridge: Cambridge University Press, 1990.);
(Mack D, Fische W, Krokotsc A, Leopold K, Hartmann R, Egge H, and
Laufs R, "The intercellular adhesin involved in biofilm
accumulation of Staphylococcus epidermidis is a linear
.beta.-1,6-linked glucosaminoglycan: purification and structural
analysis," J Bacteriol, 1996; 178: 175-183.).
Because only small amounts of the biofilm-derived EPS are normally
available for direct studies, the researchers usually use data
derived from planktonic cell cultures and extrapolate them to
biofilms. There is no conclusive evidence to support the idea of
existence of the biofilm-specific polysaccharides, and to date, all
studied polysaccharides present in various biofilms resemble
closely the corresponding polymers synthesized by planktonic cells.
It has been proposed that the increased amount of polysaccharides
in biofilm (one or more, specific for a given bacteria in any given
biofilm) can be part of a stress response in biofilm-grown
microorganisms, and bacteria form exopolysaccharides as a
by-product to release reducing equivalents accumulated in
non-optimal growth conditions (Creti R, Koch S, Fabretti F,
Baldassarri L, and Huebneri J, "Enterococcal colonization of the
gastro-intestinal tract: role of biofilm and environmental
oligosaccharides," BMC Microbiology, 2006; 6: 60 doi:
10.1186/1471-2180-6-60.); (Rinker K D, Kelly R M, "Effect of carbon
and nitrogen sources on growth dynamics and exopolysaccharide
production for the hyperthermophilic archaeon Thermococcus
litoralis and bacterium Thermotoga maritime," Biotechnol Bioeng,
2000; 69: 537-547.); (Sutherland I W, "Biofilm exopolysaccharides:
a strong and sticky framework," Microbiology, 2001; 147: 3-9.).
Other extracellular products (specific substances or by-products of
bacterial metabolism), as well as detritus, can be either released
into the biofilm from aging and lysed cells or trapped within the
biofilm matrix, and "cemented" there by mixture of
exopolysaccharides (Christensen B E, "The role of extracellular
polysaccharides in biofilms," J. Biotechnol., 1989; 10: 181-201.).
These extracellular products include small sugars (mono-,
disaccharides), polyols, proteins, glycoproteins, enzymes, lipids,
glycolipids, phospholipids, nucleic acids, and DNA (Boyd A and
Chakrabarty A M, "Role of alginate lyase in cell detachment of
Pseudomonas aeruginosa," Appl Environ Microbiol, 1994; 60:
2355-2359.); (Harz M, Rosch P, Peschke K D, Ronneberger O,
Burkhardt H, and Popp J, "Micro-Raman spectroscopic identification
of bacterial cells of the genus Staphylococcus and dependence on
their cultivation conditions," Analyst, 2005; 130: 1543-1550.);
(Nottingher I, Verrier S, Haque S, Polak J M, Hench L L,
"Spectroscopic study of human lung epithelial cells (A549) in
culture: living cells versus dead cells," Biopolymers, 2003; 72:
230-240.); (Sutherland I W, "A natural terrestrial biofilm," J Ind
Microbiol, 1996; 17: 281-283.); (Webb J S et al, "Cell death in
Pseudomonas aeruginosa biofilm development," J. Bacteriol., 2003;
185: 4585-4592.); (Weldon M K, Zhelyaskov V R, Morris M D,
"Surface-enhanced Raman spectroscopy of lipids on silver
microprobes," Appl Spectrosc, 1998; 52: 265-269.); (Yarwood J M, et
al., "Quorum sensing in Staphylococcus aureus biofilms," J.
Bacteriol., 2004; 186: 1838-1850.). It has been suggested that
extracellular DNA, released from the lysed cells, plays an
important role in supporting the biofilm structure and provides
opportunities for microorganisms to exchange the genetic material
for possible development of the biofilm-specific phenotypes
(Costerton J W, Veeh R, Shirtliff M, Pasmore M, Post C, and Ehrich
G D, "The application of biofilm science to the study and control
of chronic bacterial infections," J. Clin. Invest., 2003; 112:
1466-1477.); (Gilbert P, Maira-Litran T, McBain A J, Rickard A H,
and Whyte F W, "The physiology and collective recalcitrance of
microbial biofilm communities," Adv. Microb. Physiol., 2002; 46:
202-256.); (Osterreicher-Ravid D, Ron E Z, & Rosenberg E,
"Horizontal transfer of an exopolymer complex from one bacterial
species to another," Environ Microbiol, 2000; 2: 366-372.);
(Stoodley P, Sauer K, Davies D G, and Costerton J W, "Biofilms as
complex differentiated communities," Annu. Rev. Microbiol., 2002;
56: 187-209.); (Whitchurch C B, et al., "Extracellular DNA required
for bacterial biofilm formation," Science, 2002; 295: 1487.).
It has been proposed that in the dynamic environment of biofilm,
microorganisms use special chemical signaling molecules to
communicate (the process called quorum-sensing--QS), and the
presence of an adequate number of neighboring cells with
coordinated chemical signaling between them allow bacteria to
properly respond to changes in environmental conditions, including
insult from antimicrobials, and benefit from living in the biofilm
community. It was assumed that QS can regulate extracellular
polysaccharide production, based on the major alterations in the
extracellular matrix of laboratory-grown Pseudomonas aeruginosa
biofilm when the mutant strain was unable to produce the
N-(3-oxododecanoyl)-L-homoserine lactone signal specific for QS
(Davies D, Parsek M, Pearson J, et al., "The involvement of
cell-to-cell signals in the development of a bacterial biofilm,"
Science, 1998; 280: 295-298.); (Singh P, Schaeffer A, Parsek M, et
al., "Quorum sensing signals indicate that cystic fibrosis lungs
are infected with bacterial biofilms," Nature, 2000; 407:
762-764.). But to date, the quorum-sensing-regulated genes involved
in Pseudomonas aeruginosa biofilm matrix production have not been
identified, and the pel and/or psl genes (regulating production of
other polysaccharides PEL and PSL) have not been revealed as
quorum-sensing-regulated genes as well (Branda S S, Vik A, Friedman
L, and Kolter R, "Biofilms: the matrix revisited," Trends in
Microbiology, 2005; 13(1): 20-26.); (Whiteley M, et al.,
"Identification of genes controlled by quorum sensing in
Pseudomonas aeruginosa," Proc. Natl. Acad. Sci. U.S.A., 1999; 96:
13904-13909.). Also, the role of quorum sensing in resistance of
biofilm to antimicrobials is not clear yet; for example, the
laboratory mutants defective in quorum sensing, are unaffected in
their resistance to detergents and antibiotics (Brooun A, et al.,
"A dose-response study of antibiotic resistance in Pseudomonas
aeruginosa biofilms," Antimicrob. Agents Chemother, 2000; 44:
640-646.).
According to a classical model, any biofilm can be described as: a
non-homogenous multi-layer structure with dynamic environment;
growing in a 3-dimensional mode, with constant addition of the new
layers and detachment of the parts of the biofilm; with spatial and
temporal heterogeneity within the biofilm and variations in
bacterial growth rate; with different metabolic and genetic
activities of the microorganisms resulting in increased resistance
to antimicrobials (including antibiotics) and host defense
mechanisms (Charaklis W G, Marshall K C, "Biofilm as a basis for
interdisciplinary approach," pp. 3-15, In: Biofilms, 1990, John
Wiley and Sons, Charaklis W G. and Marshall K C. (ed.), New York,
N.Y.); (Fux C A, et al., "Review. Survival strategies of infectious
biofilms", Trends in Microbiology, January 2005; Vol. 13, No 1:
34-40.). The heterogeneity within the biofilm has been confirmed
for protein synthesis and respiratory activity, but the DNA content
remained relatively constant throughout biofilm (Wentland E J, et
al., "Spatial variations in growth rate within Klebsiella
pneumoniae colonies and biofilm," Biotechnol. Prog., 1996; 12:
316-321.); (Xu K D, et al., "Biofilm resistance to antimicrobial
agents," Microbiology, 2000; 146: 547-549.). An oxygen tension
gradient exists within biofilm with the superficial areas being
more metabolically active than the deeper areas where bacteria
adapt to decreased oxygen availability (De Beer D, Stoodley P, Roe
F, et al., "Effects of biofilm structure on oxygen distribution and
mass transport," Biotechnology Bioengineering, 1994; 43:
1131-1138.). The outer layers of biofilm are more permeable to
antimicrobials due to slow build-up of polysaccharides and other
constituents (proteins, lipids, etc.), and the inner (deeper)
layers are more dense, compressed, and less permeable. Bacteria in
the outer layers of biofilm, exposed to the bulk medium, grow
faster and can be less resistant to antimicrobials. Conversely, the
bacteria in the inner or deeper layers, located closer to the
attached surface, grow slower, adapting to decreased oxygen and
nutrients availability, and in time, can become more resistant to
antimicrobials with possible consequent emergence of
biofilm-specific antibiotic-resistant phenotype (Brown M R, et al.,
"Resistance of bacterial biofilms to antibiotics: a growth-rate
related effect?," J. Antimicrob. Chemother., 1998; 22:
777-780.).
It has been proposed that "any given cell within the biofilm will
experience a slightly different environment compared with other
cells within the same biofilm, and thus be growing at a different
rate" (Mah T C, and O' Toole G A, "Review. Mechanisms of biofilm
resistance to antimicrobial agents," Trends in Microbiology,
January 2001, 9(1): 34-39.). With continuous bacterial growth,
increased cell density triggers the general stress response in
microbial cells, as confirmed by increased production of
osmoprotectant trehalose and degrading enzyme catalase, with higher
concentration of trehalose in proximity to the pathogenic cell
colonies (Liu X, et al., "Global adaptations resulting from high
population densities in Escherichia coli cultures," J. Bacteriol.,
2000; 182: 4158-4164.). These events result in physiological
changes in biofilm, including reduced flow of solutes (nutrients)
into biofilm and diminished growth rate of bacterial microcolonies
for genotype survival (Brown M R, and Barker J, "Unexplored
reservoirs of pathogenic bacteria: protozoa and biofilms," Trends
Microbiol., 1999; 7: 46-50.); (Mah T C., and O' Toole G A, "Review:
Mechanisms of biofilm resistance to antimicrobial agents", Trends
in Microbiology, January 2001; 9(1): 34-39.).
About two decades ago, the existence of biofilm-specific phenotypes
of bacteria was an emerging idea. Such biofilm-specific phenotypes,
thought to be induced in a subpopulation of microorganisms upon
attachment to a surface, were proposed to express specific
biofilm-related genes compared with their planktonic counterparts
(Kuchma S L, and O'Toole G A, "Surface-induced and biofilm-induced
changes in gene expression," Curr. Opin. Biotechnol., 2000; 11:
429-433.). Multiple research data, based mostly upon the genetic
studies of the laboratory-constructed and laboratory-grown mutant
strains, provided supportive evidence that the biofilm-grown cells
differ from their planktonic counterparts in specific properties,
including nutrients utilization, growth rate, stress response, and
increased resistance to antimicrobial agents and the host
defenses.
Biofilm Resistance to Antimicrobial Agents
The mechanism of resistance to antimicrobial agents (including
antibiotics) in biofilm-related microorganisms is different from
plasmid, transposons, and mutations that confer innate resistance
in individual bacterial cells (Stewart P S and Costerton J W,
"Review. Antibiotic resistance of bacteria in biofilms," Lancet,
2001; 358: 135-138.); (Costerton J W, Stewart P S, and Greenberg E,
"Bacterial biofilms: a common cause of persistent infections,"
Science, 1999; 284: 1318-1322.); (Costerton J W and Stewart P S,
"Biofilms and device-related infections," In: Nataro J P, Blaser M
J, Cunningham-Rundles S., (eds.), Persistent bacterial infections.
Washington, D.C.: ASM Press, 2000; 432-439.).
Multiple research studies provided basis for various mechanisms of
biofilm resistance to antimicrobials, including: physical and/or
chemical diffusion barriers to penetration of antimicrobials and
host defense cells into the exopolymer matrix of biofilm activation
of a general stress response of the microorganisms slow growth of
the microorganisms possible emergence of a biofilm-specific
bacterial phenotype
These mechanisms can be applied to any type of biofilm, varying
with the bacteria present and the type of antimicrobials being used
(Geddes A, "Infection in the twenty-first century: Predictions and
postulates," J Antimicrob Chemother, 2000; 46: 873-878.); (Stewart
P S, "Theoretical aspects of antibiotic diffusion into microbial
biofilms," Antimicrob. Agents Chemother., 1996; 40: 2517-2522.);
(Stewart P S, "Mechanisms of antibiotic resistance in bacterial
biofilms," Int J Med Microbiol, 2002; 292: 107-113.).
Most of the biofilm-resistance mechanisms are provided by the
biofilm exopolymer matrix as the initial physical and/or chemical
barrier that can prevent, inhibit or delay penetration of
antimicrobials and host defense cells into the biofilm. The
diffusion of antimicrobials through the biofilm can be inhibited by
various means: for example, the common disinfectant chlorine is
consumed by chemical reaction within the matrix of a mixed
Klebsiella pneumoniae and Pseudomonas aeruginosa biofilm (de Beer
D, et al., "Direct measurement of chlorine penetration into
biofilms during disinfection," Appl. Environ. Microbiol., 1994; 60:
4339-4344.); antibiotic ciprofloxacin binds to the biofilm
components (Suci P A, et al., "Investigation of ciprofloxacin
penetration into Pseudomonas aeruginosa biofilms," Antimicrob
Agents Chemother, 1994; 38: 2125-2133.); Pseudomonas aeruginosa
biofilm prevents diffusion of piperacillin (Hoyle B, et al.,
"Pseudomonas aeruginosa biofilm as a diffusion barrier to
piperacillin," Antimicrob. Agents Chemother., 1992: 36:
2054-2056.); positively charged aminoglycosides bind to negatively
charged matrix polymers, such as .beta.1,4-glucosaminoglycan in
Staphylococcus epidermidis biofilm and alginate in Pseudomonas
aeruginosa biofilm (Lewis K, "Riddle of biofilm resistance,"
Antimicrob Agents Chemother., 2001; 45: 999-1007.); (Walters M C,
et al., "Contributions of antibiotic penetration, oxygen
limitation, and low metabolic activity to tolerance of Pseudomonas
aeruginosa biofilms to ciprofloxacin and tobramycin," Antimicrob.
Agents Chemother., 2003; 47: 317-323.); (Gordon C A, Hodges N A,
Marriott C, "Antibiotic interaction and diffusion through alginate
exopolysaccharide of Cystic fibrosis--derived Pseudomonas
aeruginosa," J. Antimicrob. Chemother., 1988; 22: 667-674.);
(Nichols W W, et al., "Inhibition of tobramycin diffusion by
binding to alginate," Antimicrob. Agents Chemother., 1988; 32:
518-523.); the additional matrix component colanic acid, produced
by mucoid phenotype of E. coli, supports biofilm maturation and
provides a thicker biofilm (Danese P N, et al., "Exopolysaccharide
production is required for development of Escherichia coli K-12
biofilm architecture," J. Bacteriol., 2000; 182: 3593-3596.);
penetration of antifungal agent nystatin into the mycelium of
Aspergillus fumigatus submerged in medium and covered by thin layer
of exopolymer matrix is higher than into the aerial-grown colony
covered by thick layer of extracellular matrix (Beauvais A, et al.,
"An extracellular matrix glues together the aerial-grown hyphae of
Aspergillus fumigatus," Cellular Microbiology, 2007; 9 (6):
1588-1600.); secreted IgG antibodies fail to penetrate biofilm
because of matrix binding (de Beer D, et al., "Measurement of local
diffusion coefficients in biofilms by micro-injection and confocal
microscopy," Biotechnol. Bioeng., 1997; 53: 151-158.); alginate
produced by mucoid phenotype of Pseudomonas aeruginosa protects
bacteria from phagocytosis by host leukocytes and TNF-.gamma.
activated macrophages (Bayer A S, et al., "Functional role of
mucoid exopolysaccharide (alginate) in antibiotic-induced and
polymorphonuclear leukocyte-mediated killing of Pseudomonas
aeruginosa," Infect. Immun., 1991; 59: 302-308.); (Leid J G,
Willson C J, Shirtliff M E, Hassett D J, Parsek M R, and Jeffers A
K, "The exopolysaccharide alginate protects Pseudomonas aeruginosa
biofilm bacteria from IFN-gamma-mediated macrophage killing." J
Immunol, 2005; 175: 7512-7518.).
Antimicrobials diffusion can also be inhibited or delayed by
specific active substances produced by bacteria themselves: for
example, enzyme catalase produced by Pseudomonas aeruginosa spp.
degrades hydrogen peroxide on diffusion into thick biofilm (Stewart
P S, et al., "Effect of catalase on hydrogen peroxide penetration
into Pseudomonas aeruginosa biofilms," Appl. Environ. Microbiol.,
2000; 66: 836-838.); ampicillin is unable to penetrate biofilm of
Klebsiella pneumoniae due to ampicillin-degrading enzyme
Beta-lactamase (Anderi J N, et al., "Role of antibiotic penetration
limitation in Klebsiella pneumoniae biofilm resistance to
ampicillin and ciprofloxacin," Antimicrob. Agents Chemother., 2000;
44: 1818-1824.); (Bagge N, Hentzer M, Andersen J B, Ciofu O,
Givskov M, and Hoiby N, "Dynamics and spatial distribution of
beta-lactamase expression in Pseudomonas aeruginosa biofilms,"
Antimicrob Agents Chemother, 2004; 48: 1168-1174.); extracellular
slime derived from coagulase-negative Staphylococci reduces the
effect of glycopeptide antibiotics (Konig C, et al., "Factors
compromising antibiotic activity against biofilms of Staphylococcus
epidermidis," Eur. J. Clin. Microbiol. Infect. Dis., 2001; 20:
20-26.); (Souli M and Giamarellou H., "Effects of slime produced by
clinical isolates of coagulase-negative staphylococci on activities
of various antimicrobial agents," Antimicrob. Agents Chemother.,
1998; 42: 939-941.); a PMN toxin, rhamnolipid B, produced by
Pseudomonas aeruginosa is known to kill neutrophils (Jensen P O,
Bjarnsholt T, Phipps R, Rasmussen T B, Calum H, Christoffersen L,
et al., "Rapid necrotic killing of polymorphonuclear leukocytes is
caused by quorum-sensing-controlled production of rhamnolipid by
Pseudomonas aeruginosa," Microbiology, 2007; 153: 1329-1338.).
Delayed penetration of antimicrobials into the biofilm can provide
enough time for bacteria to induce the expression of various genes
regulating the stress response and mediating resistance to
antimicrobials (Jefferson K K, Goldmann D A, and Pier G B, "Use of
confocal microscopy to analyze the rate of vancomycin penetration
through Staphylococcus aureus biofilms," Antimicrob Agents
Chemother, 2005; 49: 2467-2473.); (Anwar H, Strap J L, and
Costerton J W, "Establishment of aging biofilms: a possible
mechanism of bacterial resistance to antimicrobial therapy,"
Antimicrob Agents Chemother, 1992; 36: 1347-1351.). The central
regulator of a general stress response is the alternate
sigma-factor RpoS induced by high cell density, and the presence of
activated gene rpoS' mRNA was detected by RT-PCR in sputum from
Cystic Fibrosis patients with chronic Pseudomonas aeruginosa
biofilm infections (Foley I, et al., "General stress response
master regulator rpoS is expressed in human infection: a possible
role in chronicity," J. Antimicrob. Chemother., 1999; 43:
164-165.). Also, it has been shown that an additional sigma-factor
Alg acted in concert with RpoS to control general stress response
in laboratory grown Pseudomonas aeruginosa during biofilm formation
and maturation, and several other genes were upregulated as well,
including algC (controlling phosphomannomutase, involved in
exopolysaccharide alginate synthesis), algD, algU, and genes
controlling polyphosphokinase synthesis (Davis D G and Geesey G G,
"Regulation of the alginate biosynsthesis gene algC in Pseudomonas
aeruginosa during biofilm development in continuous culture," Appl.
Environ. Microbiol., 1995; 61: 860-867.). It has been demonstrated
that as many as 45 genes differed in expression between sessile
cells and their planktonic counterparts during the biofilm
development in laboratory settings.
Biofilm-Based Medical Conditions and Diseases
Comprehensive review of the biofilm-based human infections as well
as the biofilms on medical devices was published by Rodney M.
Donlan and J. William Costerton (Donlan R M and Costerton J W,
"Review. Biofilms: Survival mechanisms of clinically relevant
microorganisms," Clinical Microbiology Reviews, April 2002;
167-193.). Microbial biofilms are important factors in the
pathogenesis of various human chronic infections, including native
valve endocarditis (NVE), line sepsis, chronic otitis media,
chronic sinusitis and rhinosinusitis, chronic bronchitis, cystic
fibrosis pseudomonas pneumonia, chronic bacterial prostatitis,
chronic urinary tract infections (UTIs), periodontal disease,
chronic wound infections, osteomyelitis (Costerton J W, Stewart P,
Greenberg E, "Bacterial biofilms: a common cause of persistent
infections," Science, 1999; 284: 1318-1322.); (Hall-Stoodley L and
Stoodley P, "Evolving concepts in biofilm infections," Cellular
Microbiology, 2009; 11 (7): 1034-1043.). Microbial biofilms are
detected on various medical devices (prosthetic heart valves,
central venous catheters, urinary catheters, contact lenses,
tympanostomy tubes, intrauterine devices), as well as on medical
equipment (endoscopes, dialysis systems, nebulizers, dental unit
water lines), and on a variety of surfaces in hospitals and other
medical settings (Costeron J W and Stewart P S, "Biofilms and
device-related infections," In: Nataro J. P., Blaser M. J.,
Cunningham-Rundles S., eds. Persistent bacterial infections.
Washington, D.C.: ASM Press, 2000; 432-439.); (Bryers J D, "Medical
Biofilms," Biotechnology and Bioengineering, 2008; 100 (1) May 1.).
Due to their specific features, chronic biofilm-based infections
require different interventional approaches for effective treatment
(Stewart P S and Costerton J W., "Review. Antibiotic resistance of
bacteria in biofilms," Lancet, 2001; 358: 135-138.); (Donlan R M
and Costerton J W, "Review. Biofilms: Survival mechanisms of
clinically relevant microorganisms," Clinical Microbiology Reviews,
April 2002; 167-193.); (Costerton J W, Stewart P S, and Greenberg E
P, "Bacterial biofilms: a common cause of persistent infections,"
Science, 1999; 284: 1318-1322.); (Costerton J W and Stewart P S,
"Biofilms and device-related infections," In: Nataro J P, Blaser M
J, Cunningham-Rundles S, eds. Persistent bacterial infections.
Washington, D.C.: ASM Press, 2000; 432-439.); (Wolcott R D, M. D.
and Ehrlich G D, Ph.D., "Biofilms and chronic infections," JAMA,
2008, Vol. 299, No 22.); (Costerton J W, Irvin R T, "The Bacteria
Glycocalyx in Nature and Disease," Ann. Rev. Microbiol., 1981; 35:
299-324.); (Costerton J W, et al., "The application of biofilm
science to the study and control of chronic bacterial infections,"
J. Clin. Invest., 2003; 112: 1466-1477.).
Native Valve Endocarditis
The development of Native Valve Endocarditis (NVE) results from the
interaction between the endothelium of the heart (generally, of the
mitral, aortic, tricuspid, and pulmonic valves) and microorganisms
circulating in the bloodstream (Livornese L L and Korzeniowski O M,
"Pathogenesis of infective endocarditis," pp. 19-35. In: Infective
endocarditis, Kaye D. (ed.), 2-nd ed., 1992; Raven Press, New York,
N.Y.). Microorganisms usually do not adhere to intact endothelium.
There should be contributing factors that promote adherence, such
as: damaged endothelium (as in vasculitis), formation of initial
thrombotic lesions of heart valves (as in nonbacterial thrombotic
endocarditis--NBTE), accumulation of fibronectin secreted by
endothelial cells, platelets and fibroblasts in response to
vascular injury, which can simultaneously bind to fibrin, collagen,
human cells, and bacteria, specific fibronectin receptors in some
bacteria (Streptococcus sanguis, Staphylococcus aureus),
high-molecular weight dextrans produced by various Streptococci
that promote adherence to the surface of the thrombus in NBTE
(Lowrance J H, Baddour L M, and Simpson W A, "The role of
fibronectin binding on the rate model of experimental endocarditis
caused by Streptococcus sanguis," J. Clin. Investig. 86: 7-13.);
(Roberts R B, "Streptococcal endocarditis: the viridins and beta
hemolytic streptococci," pp. 191-208. In: Infective endocarditis,
Kaye D. (ed.), 2-nd ed., 1992; Raven Press, New York, N.Y.). The
most metabolically active colonies were detected on the surface of
the thrombus, forming initial biofilm there (Durack D T and Beeson
P B, "Experimental bacterial endocarditis II. Survival of bacteria
in endocardial vegetations," Br. J. Pathol., 1972, 53: 50-53.).
Clinical research of 2345 cases of NVE demonstrated a variety of
microorganisms involved: Streptococci (including Streptococcus
viridans, Streptococcus bovis), Enterococci, Pneumococci .about.in
56% of cases; Staphylococci .about.in 25% of cases
(.about.19%--Coagulase positive and .about.6%--Coagulase negative);
Gram-negative bacteria .about.in 11% of cases, and Fungi (Candida
and Aspergillus spp.) .about.in 10% of cases; all these
microorganisms gained access to the bloodstream primarily via the
oropharynx, gastrointestinal tract, and genitourinary tract (Tunkel
A R and Mandell G I, "Infecting microorganisms," pp. 85-97. In:
Infective endocarditis, Kaye D. (ed.), 2-nd ed., 1992; Raven Press,
New York, N.Y.).
Biofilm-Based Chronic Infections in the Respiratory Tract
In the upper respiratory tract, bacterial biofilms have been
demonstrated in chronic tonsillitis, chronic adenoiditis, chronic
sinusitis and chronic rhinosinusitis (CRS), chronic otitis media
(OM), and cholesteatoma. In clinical specimens from patients with
chronic and recurrent tonsillitis, both attached and aggregated
biofilm-associated bacteria were detected in mucosal epithelium of
tonsils removed for chronic tonsillitis (in 73% of cases) and in
75% of cases of tonsils removed due to hypertrophy alone (Chole R A
and Faddis B T, "Anatomical evidence of microbial biofilms in
tonsillar tissues: a possible mechanism to explain chronicity,"
Arch Otolaryngol Head Neck Surg, 2003; 129: 634-636.). Microbial
biofilms associated with epithelial lining with presence of a
carbohydrate matrix in situ were demonstrated in clinical specimens
of human adenoids removed for chronic adenoiditis (Kania R E,
Lamers G E, Vonk M J, Dorpmans E, Struik J, Tran Ba Huy P, et al.,
"Characterization of mucosal biofilms on human adenoid tissues,"
Laryngoscope, 2008; 118: 128-134.); (Nistico L, Gieseke A, Stoodley
P, Hall-Stoodley L, Kerschner J E, and Ehrlich G D, "Fluorescence
`in situ` hybridization for the detection of biofilm in the middle
ear and upper respiratory tract mucosa," Methods Mol Biol, 2009;
493: 191-213.).
Chronic Rhinosinusitis
In Chronic Rhinosinusitis (CRS), mucosal changes with different
degrees of denudation in epithelial cells result in a surface
favorable for bacterial colonization and biofilm development
(Biedlingmaier J, Trifillis A, "Comparison of CT scan and electron
microscopic findings on endoscopically harvested middle
turbinates," Otolaryngol Head Neck Surg, 1998; 118: 165-173.).
Biofilm formation, mainly with Pseudomonas aeruginosa infection,
was confirmed in patients who had surgery and continued to have
symptoms despite medical treatment (Cryer J, Schipor I, Perloff J
R, Palmer J N, "Evidence of bacterial biofilms in human chronic
sinusitis," ORL J Otolaryngol Relat Spec, 2004; 66: 155-158.). In
patients with CRS having surgery, mucosal biopsies demonstrated
different stages of the biofilm by scanning electron microscopy
(SEM) in five out of five patients, and all five patients showed
aberrant findings of the mucosal surface with various degrees of
severity: from disarrayed cilia to complete absence of cilia and
goblet cells (Ramadan H H, Sanclement J A, Thomas J G, "Chronic
rhinosinusitis and biofilms," Otolaryngol Head Neck Surg, 2005;
132: 414-417.). In most cases of CRS and Pseudomonas aeruginosa
biofilms, clinical symptoms were refractory to culture-directed
antibiotics, topical steroids, and nasal lavages, and only surgery
(mechanical debridement) resulted in significant improvement
(Ferguson B J, Stolz D B, "Demonstration of biofilm in human
bacterial chronic rhinosinusitis," Am J Rhinol, 2005; 19:
452-457.).
Chronic Otitis Media
Chronic Otitis Media (OM) involves inflammation of the middle-ear
mucoperiosteal lining and is caused by a variety of microorganisms,
including: Streptococcus pneumoniae, Haemophilus influenzae,
Moraxella catarrhalis, group A beta-hemolytic streptococci, enteric
bacteria, Staphylococcus aureus, Staphylococcus epidermidis,
Pseudomonas aeruginosa, and other organisms; mixed cultures can
also be isolated (Feigin R D, Kline M W, Hyatt S R, and Ford III K
L, "Otitis media," pp. 174-189. In: Textbook of pediatric
infectious diseases, Feigin R D and Cherry J D (ed.), 3-rd ed.,
vol. 1, 1992, W. B. Saunders Co., Philadelphia, Pa.); (Giebink G S,
Juhn S K, Weber M L, and Le C T, "The bacteriology and cytology of
chronic otitis media with effusion," Pediatric Infect. Dis., 1982;
1: 98-103.). Chronic OM as a biofilm-related infection was
demonstrated in clinical specimens and in animal models. Scanning
electron microscopy provided evidence of Haemophilus influenzae
biofilm on the middle-ear mucosal surfaces of chinchillas that had
been injected with a culture of this organism (Hayes J D, Veeh R,
Wang X, Costerton J W, Post J C, and Ehrlich G D, Abstr. 186, Am.
Soc. Microbiol. Biofilm, 2000; Conf. 2000.); (Hong W, Mason K,
Jurcisek J, Novotny L, Bakaletz L O, and Swords W E,
"Phosphorylcholine decreases early inflammation and promotes the
establishment of stable biofilm communities of nontypeable
Haemophilus influenzae strain 86-028NP in a chinchilla model of
otitis media," Infect Immun, 2007b; 75: 958-965.). Biofilm
aggregates of Streptococcus pneumoniae, Haemophilus influenzae and
Moraxella catarrhalis were detected in biopsies of the middle-ear
mucosal lining in children with chronic or recurrent OM undergoing
TT placement for treatment, but not in the middle-ear mucosal
biopsies from patients undergoing surgery for cochlear implantation
(Hall-Stoodley L, Hu F Z, Gieseke A, Nistico L, Nguyen D, Hayes J,
et al., "Direct detection of bacterial biofilms on the middle-ear
mucosa of children with chronic otitis media," JAMA, 2006; 296:
202-211.)
In chronic OM with effusion, the presence of highly viscous fluid
in the middle ear requires in many cases the implantation of
tympanostomy tubes (TT) to alleviate pressure build-up and hearing
loss. Tympanostomy tubes are subject to contamination, and biofilms
build up on their inner surfaces. The investigation of colonization
and biofilm development by Pseudomonas aeruginosa, Staphylococcus
aureus, and Staphylococcus epidermidis on various tympanostomy
tubes, provided evidence that all three organisms developed
biofilms on the Armstrong silicone and the silver oxide-coated
Armstrong-style silicone tubes; Pseudomonas aeruginosa also
developed biofilms on the fluoroplastic tubes; only the ionized
silicone tubes remained free of contamination and biofilms
(Biedlingmaier J F, Samaranayake R, and Whelan P, "Resistance to
biofilm formation on otologic implant materials," Otolaryngol Head
Neck Surg, 1998; 118: 444-451.). Silver oxide-impregnated silastic
tubes lowered the incidence of postoperative otorrhea during the
first postoperative week, possibly by preventing adherence and
colonization of selected bacteria to the tube, but had no effect on
the established infection in the middle ear (Gourin C G and Hubbell
R N, "Otorrhea after insertion of silver oxide-impregnated silastic
tympanostomy tubes," Arch. Otolaryngol Head Neck Surg, 1999; 125:
446-450.). Bacterial biofilm was also detected on a human cochlear
implant (Pawlowski K S, Wawro D, Roland P S, "Bacterial biofilm
formation on a human cochlear implant," Otol Neurotol, 2005; 26:
972-975.).
In the lower respiratory tract, microbial biofilms were associated
with chronic bronchitis, chronic obstructive pulmonary disease, and
pneumonia, especially in patients with cystic fibrosis. Scanning
electron microscopy of clinical samples (sputum, bronchiolar
lavage, lung and bronchial lining biopsies) demonstrated microbial
biofilms either attached to mucosal linings or in the form of
bacterial aggregates in mucus covering respiratory epithelium (Lam
J, Chan R, Lam K, and Costerton J W, "Production of mucoid
microcolonies by Pseudomonas aeruginosa within infected lungs in
cystic fibrosis," Infect Immun, 1980; 28: 546-556.);
(Martinez-Solano L, Macia M D, Fajardo A, Oliver A, and Martinez J
L, "Chronic Pseudomonas aeruginosa infection in chronic obstructive
pulmonary disease," Clin Infect Dis, 2008; 47: 1526-1533.);
(Starner T D, Zhang N, Kim G, Apicella M A, and McCray P B Jr,
"Haemophilus influenzae forms biofilms on airway epithelia:
implications in cystic fibrosis," Am J Respir Crit Care Med, 2006;
174: 213-220.); (Worlitzsch D, Tarran R, Ulrich M, Schwab U, Cekici
A, Meyer K C, et al., 2002, "Effects of reduced mucus oxygen
concentration in airway Pseudomonas infections of cystic fibrosis
patients," J Clin Invest, 2002; 109: 317-325.); (Yang L, Haagensen
J A, Jelsbak L, Johansen H K, Sternberg C, Hoiby N, and Molin S,
"In situ growth rates and biofilm development of Pseudomonas
aeruginosa populations in chronic lung infections," J Bacteriol,
2008; 190: 2767-2776.).
Cystic Fibrosis
Cystic fibrosis (CF), a chronic disease of the lower respiratory
system, is the most common inherited disease: 70% of patients with
CF are defective in the cystic fibrosis transmembrane conductance
regulator protein (CFTR), which functions as a chloride ion channel
protein, resulting in altered secretions in the secretory epithelia
of the respiratory tract. In CF, there is a net deficiency of
water, which hinders the upward flow of the mucus layer thus
altering mucociliary clearance. Decreased secretion and increased
absorption of electrolytes lead to dehydration and thickening of
secretions covering the respiratory mucosa (Koch C and Hoiby N,
"Pathogenesis of cystic fibrosis," Lancet, 1993; 341: 1065-1069.).
The hyperviscous mucus is thought to increase the incidence of
bacterial lung infections in CF patients. Staphylococcus aureus is
usually the first pulmonary isolate from these patients, followed
by Haemophilus influenzae. Both of these infections can be treated
effectively with antibiotics, but on persistence, they usually form
biofilm and predispose the CF-affected lung to colonization with
Pseudomonas aeruginosa (colonization rate of .about.80%) and
Burkholderia cepacia with lethal consequences (Govan J R, and
Deretic V, "Microbial pathogenesis in cystic fibrosis: mucoid
Pseudomonas aeruginosa and Burkholderia cepacia," Microbiol. Rev.,
1996; 60: 539-574.). As was demonstrated in clinical studies, both
organisms were nonmucoid during initial colonization, but on
persistence in the lungs of patients with CF they ultimately
undergo changes to mucoid phenotype within a period of time from
months to years (Koch C and Hoiby N, "Pathogenesis of cystic
fibrosis," Lancet, 1993; 341: 1065-1069.). The mucoid material,
which was shown to be a polysaccharide substance, later identified
as alginate, was transiently produced by laboratory strain of P.
aeruginosa, following adherence to the surface (Hoyle B D, Williams
L J, and Costerton J W, "Production of mucoid exopolysaccharide
during development of Pseudomonas aeruginosa biofilms," Infect.
Immun., 1993; 61: 777-780.). It has been proposed that several in
vitro conditions, such as nutrient limitation, the addition of
surfactants, and suboptimal levels of antibiotics, may result in
mucoidy due to increased production of alginate (May T B,
Shinabarger D, Maharaj R, Kato J, Chu L, DeVault J D, Roychoudhury
S, Zielinski N A, Berry A, Rothmel R K, Misra T K, and Chakrabarty
A M, "Alginate synthesis by Pseudomonas aeruginosa: a key
pathogenic factor in chronic pulmonary infections of cystic
fibrosis patients," Clin. Microbiol. Rev., 1991; 4: 191-206.).
Early antimicrobial treatment with oral ciprofloxacin and inhaled
colistin has been shown to postpone chronic infection with
Pseudomonas aeruginosa for several years (Koch C and Hoiby N,
"Pathogenesis of cystic fibrosis," Lancet, 1993; 341:
1065-1069.).
Periodontal Diseases
Periodontal diseases include infections of the supporting tissues
of teeth, ranging from mild and reversible inflammation of the gums
(gingiva) to chronic destruction of periodontal tissues (gingiva,
periodontal ligament, and alveolar bone) and exfoliation of the
teeth. The subgingival crevice (the channel between the tooth root
and the gum) is the primary site of periodontal infection and will
deepen into a periodontal pocket with the progression of the
disease (Lamont R J and Jenkinson H F, "Life below gum line:
pathogenic mechanisms of Porphyromonas gingivalis," Microbiol. Mol.
Biol. Rev., 1998; 62: 1244-1263.). Microorganisms isolated from
patients with moderate periodontal disease include Fusobacterium
nucleatum, Peptostreptococcus micros, Eubacterium timidum,
Eubacterium brachy, Lactobacillus spp., Actinomyces naeslundii,
Pseudomonas anaerobius, Eubacterium sp. strain D8, Bacteroides
intermedius, Fusobacterium spp., Selenomonas sputigena, Eubacterium
sp. strain D6, Bacteroides pneumosintes, and Haemophilus
aphrophilus, and these bacteria are not found in healthy patients
(Moore W E C, Holdeman L V, Cato E P, Smilbert R M, Burmeister J A,
and Ranney R R, "Bacteriology of moderate (chronic) periodontitis
in mature adult humans," Infect. Immun., 1993; 42: 510-515.). In
adult patients with periodontitis, subgingival plaques harbor
spirochetes and cocci, and the predominant microorganisms of active
lesions in subgingival areas include Fusobacterium nucleatum,
Wolinella recta, Bacteroides intermedius, Bacteroides forsythus,
and Bacteroides gingivalis (Porphyromonas gingivalis) (Omar A A,
Newman H N, and Osborn J, "Darkground microscopy of subgingival
plaque from the top to the bottom of the periodontal pocket," J.
Clin. Periodontol., 1990; 17: 364-370.); (Dzink J I, Socransky S S,
and Haffajee A D, "The predominant cultivable microbiota of active
and inactive lesions of destructive periodontal diseases," J. Clin.
Periodontol., 1988; 15: 316-323.).
Proteinaceous conditioning films (called acquired pellicle),
developed on the exposed surfaces of enamel almost immediately
after cleaning of the tooth surface, comprises albumin, lysozyme,
glycoproteins, phosphoproteins, lipids, and gingival crevice fluid.
Within hours of pellicle formation, single cells of primarily
gram-positive cocci and rod-shaped bacteria from the normal oral
flora colonize these surfaces, binding directly to the pellicle
through the production of extracellular glucans (Kolenbrander P E
and London J, "Adhere today, here tomorrow: oral bacterial
adherence," J. Bacteriol., 1993; 175: 3247-3252.). After several
days, actinomycetes predominate followed by co-aggregation of
various microorganisms, resulting in the development of early
biofilm with characteristic polysaccharide matrix and polymers of
salivary origin, with subsequent (within 2 to 3 weeks) formation of
the dental plaque if left undisturbed (Marsh P D, "Dental plaque,"
pp. 282-300. In: Microbial biofilms. 1995; Lappin-Scott H M and
Costerton J W (ed.), Cambridge University Press, Cambridge, United
Kingdom.). Plaque can be mineralized with calcium and phosphate
ions (called calculus or tartar) and develop more extensively in
protected areas (between the teeth, and between the tooth and gum).
With the increase of the plaque mass in these protected areas, the
beneficial buffering and antimicrobial properties of saliva
decrease, leading to dental caries or periodontal disease. Clinical
research data show that control of supragingival plaque by
professional tooth cleaning and personal hygienic efforts can
prevent gingival inflammation and adult periodontitis (Corbet E F
and Davies W I R, "The role of supragingival plaque in the control
of progressive periodontal disease," J. Clin. Periodontol., 1993;
20: 307-313.).
Chronic Bacterial Prostatitis
The prostate gland may become infected by bacteria ascended from
the urethra or by reflux of infected urine into the prostatic ducts
emptying into the posterior urethra (Domingue G J and Hellstrom W J
G, "Prostatitis," Clin. Microbiol. Rev., 1998; 11: 604-613.). If
bacteria were not eradicated with antibiotic therapy at the early
stage of infection, they continue to persist and can form sporadic
microcolonies and biofilms that adhere to the epithelial cells of
the prostatic duct system, resulting in chronic bacterial
prostatitis. The microorganisms involved in this process include:
E. coli (most common isolate), Klebsiella, Enterobacteria, Proteus,
Serratia, Pseudomonas aeruginosa, Enterococcus fecalis, Bacteroides
spp., Gardnerella spp., Corynebacterium spp., and
Coagulase-negative Staphylococci (CONS) (Nickel J C, Costerton J W,
McLean R J C, and Olson M, "Bacterial biofilms: influence on the
pathogenesis, diagnosis, and treatment of the urinary tract
infections," J. Antimicrob. Chemother., 1994; 33 (Suppl. A):
31-41.). The biopsies from patients with chronic bacterial
prostatitis examined by either scanning electron microscopy or
transmission electron microscopy, demonstrated bacteria present in
glycocalyx-encasted microcolonies, firmly adherent to the ductal
and acinar mucosal layers (Nickel J C and Costerton J W, "Bacterial
localization in antibiotic-refractory chronic bacterial
prostatitis," Prostate, 1993; 23: 107-114.). Sporadic microcolonies
of CoNS in the intraductal space have been shown to be enveloped in
a dehydrated slime matrix (Nickel J C and Costerton J W,
"Coagulase-negative staphylococcus in chronic prostatitis," J.
Urol., 1992; 147: 398-401.). Treatment failures are common in
chronic bacterial prostatitis due to the local environment and
biofilm formation, with changes in bacterial metabolism and
possible development of resistance to antimicrobials. In order to
increase the effectiveness of the antimicrobial treatment, it has
been proposed to deliver higher antibiotic concentrations directly
to the biofilm within the prostatic ducts (Nickel J C, Costerton J
W, Mclean R J C, and Olson M, "Bacterial biofilms: influence on the
pathogenesis, diagnosis, and treatment of the urinary tract
infections," J. Antimicrob. Chemother., 1994; 33 (Suppl. A):
31-41.).
Biofilms on Medical Devices
Over the last 20 years, biofilms on various medical devices,
including prosthetic heart valves, central venous catheters,
urinary (Foley) catheters, contact lenses, intrauterine devices,
and dental unit water lines, have been studied using viable
bacterial culture techniques and scanning electron microscopy, and
for certain devices (contact lenses and urinary catheters)
additional evaluation of susceptibility of various materials to
bacterial adhesion and biofilm formation have also been implemented
(Costerton J W, Stewart P S, and Greenberg E P, "Bacterial
biofilms: a common cause of persistent infections," Science, 1999;
284: 1318-1322.); (Donlan R M and Costerton J W, "Review. Biofilms:
Survival mechanisms of clinically relevant microorganisms,"
Clinical Microbiology Reviews, April 2002; 167-193.).
Prosthetic Heart Valves
Prosthetic valve endocarditis (PVE) is a microbial infection of the
valve and surrounding tissues of the heart, ranging between 0.5%
and 4%, and is similar for both types of valves currently
used--mechanical valves and bioprostheses (Douglas J L and Cobbs C
G, "Prosthetic valve endocarditis," pp. 375-396. In: Infective
endocarditis, Kaye D. (ed.), 2-nd ed., 1992; Raven Press LTD., New
York, N.Y.). Tissue damage resulting from surgical implantation of
the prosthetic valve, leads to accumulation of platelets and fibrin
at the suture site and on the device, providing a favorable
environment for bacterial colonization and biofilm development. PVE
is predominantly caused by microbial colonization of the sewing
cuff fabric. The microorganisms commonly invade the valve annulus,
potentially promoting separation between the valve and the tissue
resulting in leakage. Infectious microorganisms involved in PVE
include Staphylococcus epidermidis (at the early stages), followed
by Streptococci, CONS, Enterococci, Staphylococcus aureus,
gram-negative Coccobacilli, fungi, and Streptococcus viridans spp.
(the most common microorganism isolated during late PVE) (Hancock E
W, "Artificial valve disease," pp. 1539-1545. In: The heart
arteries and veins; Schlant R C, Alexander R W, O'Rourke R A,
Roberts R, and Sonnenblick E H (ed.), 8-th ed., 1994; vol. 2.
McGraw-Hill, Inc., New York, N.Y.); (Illingworth B L, Twenden K,
Schroeder R F, and Cameron J D, "In vivo efficacy of silver-coated
(silzone) infection-resistant polyester fabric against a biofilm
producing bacteria, Staphylococcus epidermidis, J. Heart Valve
Dis., 1998; 7: 524-530.); (Karchmer A W and Gibbons G W,
"Infections of prosthetic heart valves and vascular grafts," pp.
213-249. In: Infections associated with indwelling medical devices;
Bisno A L and Waldovogel F A (ed.), 1994, 2-nd ed. American Society
for Microbiology, Washington, D.C.).
Central Venous Catheters
For Central Venous Catheters (CVCs), the device-related infection
rate is 3% to 5%. Infectious biofilms are universally present on
CVCs and can be associated with either the outside surface of the
catheter or the inner lumen. Colonization and biofilm formation may
occur within 3 days of catheterization. Short-term catheters (in
place for less than 10 days) usually have more extensive biofilm
formation on the external surfaces, and long-term catheters (up to
30 days) have more extensive biofilm on the internal lumen. (Raad I
I, Costerton J W, Sabharwal, Sacilowski U M, Anaissie W, and Bodey
G P, "Ultrastructural analysis of indwelling vascular catheters: a
quantitative relationship between luminal colonization and duration
of placement," J. Infect. Dis., 1993; 168: 400-407.). Colonizing
microorganisms originate either from the skin insertion site,
migrating along the external surface of the device, or from the
hub, due to manipulation by health care workers, migrating along
the inner lumen (Elliott T S J, Moss H A, Tebbs S E, Wilson I C,
Bonser R S, Graham T R, Burke L P, and Faroqui M H, "Novel approach
to investigate a source of microbial contamination of central
venous catheters," Eur. J. Clin. Microbiol. Infect. Dis., 1997; 16:
210-213.). Because the device is in direct contact with the
bloodstream, the surface becomes coated with platelets, plasma and
tissue proteins such as albumin, fibrinogen, fibronectin, and
laminin, forming conditioning films to which the bacteria are
adherent: Staphylococcus aureus adheres to fibronectin, fibrinogen,
and laminin, and Staphylococcus epidermidis adheres only to
fibronectin. Organisms colonizing CVCs include CONS, Staphylococcus
aureus, Pseudomonas aeruginosa, Klebsiella pneumoniae, Enterococcus
fecalis, and Candida albicans (Elliott T S J, Moss H A, Tebbs S E,
Wilson I C, Bonser R S, Graham T R, Burke L P, and Faroqui M H,
"Novel approach to investigate a source of microbial contamination
of central venous catheters," Eur. J. Clin. Microbiol. Infect.
Dis., 1997; 16: 210-213.).
Urinary Catheters
Urinary catheters are subject to bacterial contamination regardless
of the types of the catheter systems. In open systems, the catheter
draining into an open collection container becomes contaminated
quickly, and patients commonly develop Urinary Tract Infection
(UTI) within 3 to 4 days. In closed systems, when the catheter
empties in a securely fastened plastic collecting bag, the urine
from the patient can remain sterile for 10 to 14 days in
approximately half the patients (Kaye D and Hessen T, "Infections
associated with foreign bodies in the urinary tract," pp. 291-307.
In: Infections associated with indwelling medical devices; Bisno A
L and Waldovogel F A (ed.), 1994; 2-nd ed., American Society for
Microbiology, Washington, D.C.). Regardless of the type of the
system, with short-term catheterization (up to 7 days), 10% to 50%
of patients develop UTI, and with long-term catheterization (28
days and longer) essentially all patients develop UTI (Stickler D
J, "Bacterial biofilms and the encrustation of urethral catheters,"
Biofouling, 1996; 94: 293-305.). The risk of catheter-associated
UTI increases by approximately 10% for each day the catheter is in
place. Initially, catheters are colonized by a single
microorganism, such as Staphylococcus epidermidis, Enterococcus
fecalis, E. coli, Proteus mirabilis. Later, the number and
diversity of bacteria increase, with mixed communities containing
Providencia stuartii, Pseudomonas aeruginosa, Proteus mirabilis,
Klebsiella pneumoniae, Morganella morganii, Acinetobacter
calcoaceticus, and Enterobacter aerogenes (McLean R J C, Nickel J
C, and Olson M E, "Biofilm associated urinary tract infections,"
pp. 261-273. In: Microbial biofilms; 1995, Lappin-Scott H M and
Costerton J W (ed.), Cambridge University Press, Cambridge, United
Kingdom.).
Both in vivo and in vitro studies by scanning electron microscopy
and transmission electron microscopy provide evidence for biofilm
formation on catheters. The thickness of biofilm on silicone and
silicone-coated Foley catheters from patients undergoing long-term
catheterization ranges from 200 .mu.m to 500 .mu.m, with the
thickest biofilms formed by E. coli and Klebsiella pneumoniae (up
to 490 .mu.m). The thinnest biofilms were formed by Morganella
morganii and diphtheroids (the average .about.10 .mu.m), and these
biofilms were also patchy (Ganderton L, Chawla J, Winters C,
Wimpenny J, and Stickler D, "Scanning electron microscopy of
bacterial biofilms on indwelling bladder catheters," Eur. J. Clin.
Microbiol. Infect. Dis., 1992; 11: 789-796.).
Urinary catheter biofilms are unique, because certain
microorganisms produce enzyme urease which hydrolyzes the urea of
the urine to form free ammonia, thus raising the local pH and
allowing precipitation of minerals hydroxyapatite (calcium
phosphate) and struvite (magnesium ammonium phosphate). These
minerals become deposited in the catheter biofilms, forming a
mineral encrustation which can completely block a urinary catheter
within 3 to 5 days (Tunney M M, Jones D S, and Gorman S P, "Biofilm
and biofilm-related encrustations of urinary tract devices,"
Methods Enzymol., 1999; 310: 558-566.). The primary
urease-producing organisms in urinary catheters are Proteus
mirabilis, Morganella morganii, Pseudomonas aeruginosa, Klebsiella
pneumoniae, and Proteus vulgaris. Mineral encrustations were
observed only in catheters containing these bacteria Stickler D,
Morris N, Moreno M C, and Sabbuba N, "Studies on the formation of
crystalline bacterial biofilms on urethral catheters," Eur. J.
Clin. Microbiol. Infect. Dis., 1998; 17: 649-652.); (Stickler D,
Ganderton L, King J, Nettleton J, and Winters C, "Proteus mirabilis
biofilms and the encrustation of urethral catheters," Urol. Res.,
1993; 21: 407-411.).
Contact Lenses
Bacteria adhere readily to both types of contact lenses: soft
contact lenses (made of either hydrogel or silicone) and hard
contact lenses constructed of polymethylmethacrylate. Initial
adhesion of Pseudomonas aeruginosa to hydrogel contact lenses,
resulted within 2 hours in biofilm formation with characteristic
extracellular matrix polymers observed by transmission electron
microscopy and ruthenium red staining (Miller M J and Ahearn G,
"Adherence of Pseudomonas aeruginosa to hydrophilic contact lenses
and other substrata," J. Clin. Microbiol., 1987; 25: 1392-1397.).
The degree of attachment depended on various factors, including the
nature of the substrate, pH, electrolyte concentration, ionic
charge of the polymer, and bacterial strain tested.
Organisms that have been shown to adhere to contact lenses include:
Pseudomonas aeruginosa, Staphylococcus aureus, Staphylococcus
epidermidis, Serratia spp., E. coli, Proteus spp., and Candida spp.
(Dart J K G, "Contact lens and prosthesis infections," pp. 1-30.
In: Duane's foundations of clinical ophthalmology; Tasman W and
Jaeger E A (ed.), 1996; Lippincott-Raven, Philadelphia, Pa.). An
established biofilm was detected on the lens removed from a patient
with P. aeruginosa keratitis, as well as from the patients with
clinical diagnosis of microbial keratitis, in several cases
containing multiple species of bacteria or bacteria and fungi
(Stapleton F and Dart J, "Pseudomonas keratitis associated with
biofilm formation on a disposable soft contact lens," Br. J.
Ophthalmol., 1995; 79: 864-865.); (McLaughlin-Borlace L, Stapleton
F, Matheson M, and Dart J K G, "Bacterial biofilm on contact lenses
and lens storage cases in wearers with microbial keratitis," J.
Appl. Microbiol., 1998; 84: 827-838.).
The lens case has been implicated as the primary source of
microorganisms for contaminated lenses and lens disinfectant
solutions, with contaminated storage cases in 80% of asymptomatic
lens users (McLaughlin-Borlace L, Stapleton F, Matheson M, and Dart
J K G, "Bacterial biofilm on contact lenses and lens storage cases
in wearers with microbial keratitis," J. Appl. Microbiol., 1998;
84: 827-838.). Also, the identical organisms were isolated from the
lens cases and the corneas of infected patients. Additionally,
protozoan Acanthamoeba has been shown to be a component of these
biofilms (Dart J K G, "Contact lens and prosthesis infections," pp.
1-30. In: Duane's foundations of clinical ophthalmology; Tasman W
and Jaeger E A (ed.), 1996; Lippincott-Raven, Philadelphia, Pa.);
(McLaughlin-Borlace L, Stapleton F, Matheson M, and Dart J K G,
"Bacterial biofilm on contact lenses and lens storage cases in
wearers with microbial keratitis," J. Appl. Microbiol., 1998; 84:
827-838.).
Dental Unit Water Lines
Dental procedures may expose both patients and dental professionals
to opportunistic and pathogenic organisms originating from various
components of the dental unit. Small-bore flexible plastic tubing
supplies water (municipal or from separate reservoirs containing
distilled, filtered, or sterile water) to different hand pieces
(air-water syringe, the ultrasonic scaler, the high-speed hand
piece), and elevated bacterial counts were detected in all these
systems (Barbeau J, Tanguay R, Faucher E, Avezard C, Trudel L, Cote
L, and Prevost A P, "Multiparametric analysis of waterline
contamination in dental units," Appl. Environ. Microbiol., 1996;
62: 3954-3959.); (Furuhashi M and Miyamae T, "Prevention of
bacterial contamination of water in dental units," J. Hosp.
Infect., 1985; 6: 81-88.); (Mayo J A, Oertling K M, and Andrieu S
C, "Bacterial biofilm: a source of contamination in dental
air-water syringes," Clin. Prev. Dent., 1990; 12: 13-20.);
(Williams H N, Kelley J, Folineo D, Williams G C, Hawley C L, and
Sibiski J, "Assessing microbial contamination in clean water dental
units and compliance with disinfection protocol," JADA, 1994; 125:
1205-1211.).
Organisms generally isolated from dental water units include
Pseudomonas spp., Flavobacterium spp., Acinetobacter spp.,
Moraxella spp., Achromobacter spp., Methylobacterium spp.,
Rhodotorula spp., hyphomycetes (Cladosporium spp., Aspergillus
spp., and Penicillium spp.), Bacillus spp., Streptococcus spp.,
CONS, Micrococcus spp., Corynebacterium spp., and Legionella
pneumophila (Tall B D, Williams H N, George K S, Gray R T, and
Walch M, "Bacterial succession within a biofilm in water supply
lines of dental air-water syringes," Can. J. Microbiol., 1995; 41:
647-654.); (Whitehouse R L S, Peters E, Lizotte J, and Lilge C,
"Influence of biofilms on microbial contamination in dental unit
water," J. Dent., 1991; 19: 290-295.); (Mills S E P, Lauderdale W,
and Mayhew R B, "Reduction of microbial contamination in dental
units with povidone-iodine 10%," JADA, 1986; 113: 280-284.); (Atlas
R M, Williams J F, and Huntington M K, "Legionella contamination of
dental-unit waters," Appl. Environ. Microbiol., 1995; 61:
1208-1213.); (Callacombe S J and Fernandes L L, "Detecting
Legionella pneumophila in water systems: a comparison of various
dental units," JADA, 1995; 126: 603-608.); (Pankhurst C L,
Philpott-Howard J N, Hewitt J H, and Casewell M W, "The efficacy of
chlorination and filtration in the control and eradication of
Legionella from dental chair water systems," J. Hosp. Infect.,
1990; 16: 9-18.). The variety of microorganisms observed, were
embedded in an apparent polysaccharide matrix (Whitehouse R L S,
Peters E, Lizotte J, and Lilge C, "Influence of biofilms on
microbial contamination in dental unit water," J. Dent., 1991; 19:
290-295.). Also, amebic trophozoites and cysts, and nematodes (in
one biofilm sample) were also observed (Santiago J I, Huntington M
K, Johnston A M, Quinn R S, and Williams J F, "Microbial
contamination of dental unit waterlines: short- and long-term
effects of flushing," Gen. Dent., 1994; 42: 528-535.). A positive
correlation was found between biofilm and water counts, and by 180
days of exposure, a thick, multiple layer of extracellular
polymeric substances covered the entire surface of the dental unit
water line (Tall B D, Williams H N, George K S, Gray R T, and Walch
M, "Bacterial succession within a biofilm in water supply lines of
dental air-water syringes," Can. J. Microbiol., 1995; 41:
647-654.). Biofilms containing extensive extracellular polymer
matrix and both mixed skin flora and aquatic bacteria, were also
detected on the inner lumen of saliva ejectors (Barbeau J, ten
Bocum L, Gauthier C, and Prevost A P, "Cross contamination
potential of saliva ejectors used in dentistry," J. Hosp. Infect.,
1998; 40: 303-311.).
Methods of Treating Biofilms and Biofilm-Based Infections
Many biofilm control strategies have been proposed, applied mostly
to biofilm formed on various medical devices, including long term
antibiotics for patients using these devices, various
antimicrobials to cover the surfaces of devices, various polymer
materials, ultrasound, and low-strength electrical fields along
with disinfectants.
For biofilm-based infections in the human body, a few approaches
aimed to either eradicate or penetrate the extracellular polymeric
substances have been offered: for example, a mixture of enzymes was
effective in eradicating laboratory-grown biofilms of several
different organisms (Johansen C P, Falholt P, and Gram L,
"Enzymatic removal and disinfection of bacterial biofilm," Appl.
Environ. Microbiol., 1997; 63: 3724-3728.). Another more precise
approach was identifying the polysaccharides for a specific
organism in the biofilm and treating the biofilm with that enzyme:
for example, the specific enzyme alginate lyase allowed more
effective diffusion of gentamycin and tobramycin through alginate,
the biofilm polysaccharide of mucoid Pseudomonas aeruginosa (Hatch
R A, and Schiller N L, "Alginate lyase promotes diffusion of
aminoglycosides through the extracellular polysaccharide of mucoid
Pseudomonas aeruginosa," Antimicrob. Agents Chemother., 1998; 42:
974-977.). In addition, for the management of biofilm infections,
various antibiotics have been examined extensively in vitro and in
vivo, including aminoglycosides, fluoroquinolones, macrolides, as
well as the latest protein synthesis inhibitors (Linezolid and
Quinupristin) clinically available and appear promising for
treatment of in vivo biofilm infections (In: Biofilms, Infection,
and Antimicrobial Therapy; Edited by Pace J L, Rupp M E, and Finch
R G; Boca Raton, Fla.: CRC Press, 2006. Chapter 18, page 360.).
A review of recent patent literature summarizes citations under six
categories of current treatment approaches: 1) antibiotics and
small molecule inhibitors of new and established biofilms, 2)
quorum sensing and signaling molecules inhibitors, 3) surface
coating substances for inhibition of biofilm formation, 4)
antibodies and vaccines for infectious biofilm treatment, 5)
enzymes for degrading biofilms, and 6) bacteriophage treatment of
infectious biofilms (Lynch A S and Abbanat D, "New antibiotic
agents and approaches to treat biofilm-associated infections,"
Expert Opin. Ther. Patents, 2010; 20(10): 1373-1387.).
Additional approaches involve the use of various natural substances
and combined technologies. For example, naturally occurring
impediments to biofilm adhesion have been proposed such as,
oral-ficin, a cysteine protease derived from the Ficus glabrata
tree, which prevents biofilm-forming bacteria from adhering to
surfaces (Potera C, "A Potpourri of Probing and Treating Biofilms
of the Oral Cavity," Microbe Magazine, October 2009.). The ability
of honey to prevent quorum sensing and thereby interfere with the
formation or maintenance of biofilms suggests it can be a candidate
substance for the management of infected wounds ("The role of
biofilm in wounds," a thesis submitted to the University of Wales,
Cardiff, UK, in candidature for Ph.D. by Okhiria O A, May 2010,
Chapter 5: Antimicrobial effect of honey on biofilm and quorum
sensing: 190-234.).
An example of the use of combined technologies is the treatment of
biofilm infections on implants using ultrasound in concert with
antibiotics (Carmen J C, Roeder B L, Nelson J L, Robison Ogilvie R
L, Robison R A, Schaalje G B, and Pitt W G, "Treatment of Biofilm
Infections on Implants with Low-frequency Ultrasound and
Antibiotics," Am J Infect Control. 2005, March; 33(2): 78-82.).
Methods of Addressing Biofilm Contamination of Medical
Equipment
Bacterial and fungal biofilms develop on the various types of
medical equipment. This includes medical diagnostic devices, such
as: stethoscopes, colposcopes, nasopharyngoscopes, angiography
catheters, endoscopes, angioplasty balloon catheters; and various
permanent, semi-permanent, and temporary indwelling devices, such
as: contact lenses, intrauterine devices, dental implants, urinary
tract prostheses and catheters, peritoneal dialysis catheters,
indwelling catheters for hemodialysis and for chronic
administration of chemotherapeutic agents (Hickman catheters),
cardiac implants (pacemakers, prosthetic heart valves, ventricular
assisting devices--VAD), synthetic vascular grafts and stents,
prostheses, internal fixation devices, percutaneous sutures,
tracheal and ventilator tubing, dispensing devices such as
nebulizers, and cleaning devices such as sterilizers. Summarized
herein are the current methods employed to diminish the presence of
microbial biofilms and associated pathogens on medical
equipment.
Implants
Biofilm infections associated with indwelling medical devices and
implants are difficult to resolve using conventional antibiotics.
Antibiotic treatment requires lengthy periods of administration,
with combined antibiotics at high dose, or the temporary surgical
removal of the device or associated tissue. Newer developments,
aimed at interfering with the colonization process comprise, for
example, new biomaterials, the co-application of acoustic energy or
low-voltage electric currents with antibiotics and the development
of specific anti-biofilm agents (Jass J, Surman S, and Walker J T,
"Medical biofilms: detection, prevention, and control," Vol. 2.,
John Wiley, 2003: 261.).
Central Venous Catheters
Several studies have examined the effect of various types of
antimicrobial treatment in controlling biofilm formation on venous
catheters. The methods and materials used include adding
disinfectant to physiological flush of catheters for elimination of
microbial colonization (Freeman R, Gould F K. "Infection and
intravascular catheters," [letter]. J. Antimicrob. Chemother.,
1985; 15: 258.), impregnation of catheters with polyantimicrobials
(Darouiche R O et al., "A comparison of two
antimicrobial-impregnated central venous catheters," N Engl J Med,
1999; 340: 1-8.), coating of catheters with surfactants to bond
antibiotics to catheter surfaces (Kamal G. D., Pfaller M. A., Rempe
L. E., Jebson P. J. R., "Reduced intravascular catheter infection
by antibiotic bonding. A prospective, randomized, controlled
trial", JAMA, 1991; 265: 2364-2368.), and the use of an attachable
subcutaneous cuff containing silver ions inserted after local
application of polyantibiotic (Flowers R. H., Schwenzer K. J.,
Kopel R. F., Fisch M. J., Tucker S. I., Farr B. M., "Efficacy of an
attachable subcutaneous cuff for the prevention of intravascular
catheter-related infection", JAMA, 1989; 261: 878-883.).
Prosthetic Heart Valves
The pathogenesis of infection associated with implanted heart
valves is related to the interface between the valve and
surrounding tissue. Specifically, because implantation of a
mechanical heart valve causes tissue damage at the site of its
installation, microorganisms have an increased tendency to colonize
such locations (Donlan R M, "Biofilms and Device-Associated
Infections," Emerging Infectious Diseases Journal, March-April
2001; Vol. 7, No. 2: 277-281.). Hence, biofilms resulting from such
infections tend to favor development on the tissue surrounding the
implant or the sewing cuff fabric used to attach the device to the
tissue. Silver coating of the sewing cuff has been found to reduce
such infections (Illingworth B L, Tweden K, Schroeder R F, Cameron
J D, "In vivo efficacy of silver-coated (Silzone)
infection-resistant polyester fabric against a biofilm-producing
bacteria, Staphylococcus epidermidis", J Heart Valve Dis 1998; 7:
524. Abstract); (Carrel T, Nguyen T, Kipfer B, Althaus U,
"Definitive cure of recurrent prosthetic endocarditis using
silver-coated St. Jude medical heart valves: a preliminary case
report," J Heart Valve Dis., 1998; 7: 531. Abstract.).
Urinary Catheters
Conventional approaches to the treatment of urinary catheter
biofilms include: the use of antimicrobial ointments and
lubricants, instillation or irrigation of the bladder with
antimicrobials, use of the collection bags containing antimicrobial
agents, catheter impregnation with antimicrobial agents, and the
use of systemic antibiotics (Kaye D, Hessen M T, "Infections
associated with foreign bodies in the urinary tract," In: Bisno A.
L., Waldovogel F. A., editors. Infections associated with
indwelling medical devices. 2nd ed. Washington: American Society
for Microbiology; 1994; pp. 291-307.). Such approaches have been
found to have limited efficacy, although silver impregnation of
catheters has been found to delay onset of bacteriuria (Donlan R M,
"Biofilms and Device-Associated Infections," Emerging Infectious
Diseases Journal, March-April 2001; Vol. 7, No. 2: 277-281.). From
various materials used for catheter construction, silicone
catheters obstruct less often than latex, Teflon, or
silicone-coated latex in patients prone to catheter encrustation
(Sedor J and Mulholland S G, "Hospital-acquired urinary tract
infections associated with indwelling catheter," Urol. Clin. N.
Am., 1999; 26: 821-828.).
A new product, the UroShield.TM. System, produced by NanoVibronix
uses low cost disposable ultrasonic actuators which energize all
surfaces of the catheter thereby interfering with the attachment of
bacteria, the initial step in biofilm formation (Nagy K, Koves B,
Jickel M, Tenke P, "The effectiveness of acoustic energy induced by
UroShield device in the prevention of bacteriuria and the reduction
of patient's complaints related to long-term indwelling urinary
catheters," Poster presentation at 26th Annual Congress of the
European Association of Urology (EAU); Vienna, March 2011: No. 483.
Abstract.).
Dialysis Systems
The development of biofilms throughout hemodialysis systems has
been substantiated. In fact, some cases have been suspicious for
the outbreak of infection within dialysis centers. Furthermore, the
endotoxins and other cytokines in these biofilms can cross the
dialysis membrane and trigger the inflammatory response in the
patients (Vincent F C, Tibi A R, and Darbord J C. "A bacterial
biofilm in a hemodialysis system. Assessment of disinfection and
crossing of endotoxins," ASAIO Trans., 1989; 35: 310-313.). In a
study specific to the removal of biofilms from dialysis tubing, the
efficacy of 21 different decontamination procedures was ascertained
with the most effective treatment determined to be an acid
pre-treatment, followed by use of a concentrated bleach solution;
treatments performed at high temperature did not improve the
removal of biofilm (Marion-Ferey K, et al., "Biofilm removal from
silicone tubing: an assessment of the efficacy of dialysis machine
decontamination procedures using an in vitro model," Journal of
Hospital Infection, 2003; 53(1): 64-71.).
Given the challenge of removing biofilms from the in-place water
systems found in clinical environments, a multi-step cleaning
(removal of organic material), descaling (removal of inorganic
material), and disinfection (removal of microorganisms) process is
suggested. The most common current protocols include the following:
a) citric acid followed by bleach, b) bleach alone, c) peracetic
acid with acetic acid and hydrogen peroxide (PAA), d) citric acid
followed by autoclaving, e) citric acid at elevated temperature, f)
glycolic acid at elevated temperature, g) hot water, and h) citric
acid followed by PAA. All of these disinfection protocols appear to
be highly efficient with respect to microbial killing, but were
inefficient in reducing the amount of biofilm on affected
surfaces.
No treatment thus far has shown complete biofilm removal (and
consequently endotoxins) from silicone surfaces. Descaling by
itself is inadequate, even at high temperature. Bleach appears to
be a relatively good solitary agent for biofilm removal.
Additionally, UV irradiation has been shown to have limited impact
on biofilms; and ozone has demonstrated a higher removal efficacy,
but limited biofilm killing. It has been postulated that
destruction of both the bacteria and associated endotoxins may be
possible if super-oxidative concentrations can be achieved ("The
Role of Biofilms in Device-Related Infections," Ed. By Shirtliff M
and Leid J G; Springer-Verlag, Berlin, 2009.).
Endoscopes
In a comparative study of the efficiency of numerous detergents to
remove endoscope biofilms, it was determined that "many commonly
used enzymatic cleaners fail to reduce the viable bacterial load or
remove the bacterial EPS" (Vickery K, Pajkos A, and Cossart Y,"
Removal of biofilm from endoscopes: evaluation of detergent
efficiency, "Am J Infect Control. 2004, May; 32(3): 170-176.). Only
one cleaner containing no enzymes (produced by Whiteley Medical,
Sydney, Australia) significantly reduced bacterial viability and
residual bacterial exopolysaccharide matrix.
Noteworthy is U.S. Pat. No. 6,855,678, in which it is disclosed
that through the use of scanning electron microscopy, it has been
observed that biofilm consists of a number of layers and most
importantly, there exists a thin layer of biofilm which is adjacent
and attaches tightly to the surface of medical apparatus. The
treatment formulation advocated herein includes in combination
surfactants, solvents, co-solvents, nitrogen containing biocide,
and organic chelating agents. This composition provides a simple
non-corrosive, near neutral chemical detergent product that
reliably cleans and disinfects endoscopes and other-medical
apparatus. The hypothesized method of action is that a) the solvent
and co-solvent (example solvents include low molecular weight polar
water soluble solvents such as primary and secondary alcohols,
glycols, esters, ketones, aromatic alcohols, and cyclic nitrogen
solvents containing 8 or less carbon atoms, example co-solvents
include low molecular weight amine, amide, and methyl and ethyl
derivatives of amides) act to swell the biofilm, b) the organic
chelating agent in combination with the surfactant increases the
ability of the nitrogen containing biocide to penetrate the
biofilm, and c) the organic chelating agent in combination with the
nitrogen containing biocide act to work synergistically to dislodge
the biofilm and/or kill the microorganisms therein.
Contact Lenses
Various cleaning solutions were tested against bacterial biofilms
on contact lens storage cases, including quaternary ammonium
compounds, chlorhexidine gluconate, and hydrogen peroxide 3%.
Hydrogen peroxide 3% was most effective in inactivating 24 hr-old
biofilms formed by Pseudomonas aeruginosa, Staphylococcus
epidermidis, and Streptococcus pyogenes. Biofilm of Candida
albicans was highly resistant to all of these treatments, and
Serratia marcescens could grow in chlorhexidine disinfectant
solutions (Wilson L A., Sawant A D, and Ahearn D G, "Comparative
efficacies of soft contact lens disinfectant solutions against
microbial films in lens cases," Arch. Ophthalmol., 1991; 109:
1155-1157.); (Gandhi P A, Sawant A D, Wilson L A, and Ahearn D G,
"Adaptation and growth of Serratia marcescens in contact lens
disinfectant solutions containing chlorhexidine gluconate," Appl.
Environ. Microbiol., 1993; 59: 183-188.). It has been found that
sodium salicylate decreased initial bacterial adherence to lenses
and lens cases (Farber B F, His-Chia H, Donnenfield E D, Perry H D,
Epstein A, and Wolff A, "A novel antibiofilm technology for contact
lens solutions," Ophthalmology, 1995; 102: 831-836.).
Dental Unit Water Lines
Dental unit water lines are ideal for colonization with aquatic
bacteria and biofilm formation due to their small diameter, very
high surface-to-volume ratio, and relatively low flow rates.
Currently used flushing as treatment for reducing planktonic
bacterial load that originates from the tubing biofilm, does not
provide sufficient results, and flushing alone is ineffective
(Santiago JI, Huntington M K, Johnston A M, Quinn R S, and Williams
J F, "Microbial contamination of dental unit waterlines: short- and
long-term effects of flushing," Gen. Dent., 1994; 42: 528-535.).
Added povidone-iodine reduced contamination between 4 and 5 log
fewer bacteria per ml initially, but the levels returned to
pretreatment within 22 days (Mills S E, Lauderdale P W, and Mayhew
R B, "Reduction of microbial contamination in dental units with
povidone-iodine 10%," JADA, 1986; 113: 280-284.). Treatment with
0.5 to 1 ppm free chlorine for 10 min. each day reduced normal
bacterial counts by 2 logs from pretreatment levels, but the counts
increased again after chlorination was discontinued (Feigin R D and
Henriksen K, "Methods of disinfection of the water system of dental
units by water chlorination," J. Dent. Res., 1988; 67: 1499-1504.).
Chlorination with bleach (1:10 solution) of water systems already
contaminated with bacterial biofilms was ineffective in removing
them (Murdoch-Kinch C A, Andrews N A, Atwan S, Jude R, Gleason M J,
and Molinari J A, "Comparison of dental water quality management
procedures," JADA, 1997; 128: 1235-1243.).
Biofilms in Industrial Applications (Pipelines, Marine Biofouling,
Food Sanitation, and HVAC)
Industrial systems suffer a number of deleterious effects due to
the presence of biofilms. For heating and cooling systems, as well
as oil, water, and gas distributions systems, these effects include
flow restrictions in pipelines, flow contamination, and corrosion.
For marine systems such as ships, biofouling of hulls can lead to
tremendous loss of ship fuel efficiency owing to increased drag of
the hull.
Current Approaches for Treating Biofilms in Water, Oil and Gas
Distribution Systems
In industrial systems for the distribution of water, oil, and gas,
biofilms can form heavy biomass that can reduce the effective
diameter of a pipe or other conduit at a particular point or
increase friction along the flow path in the conduit. This
increases resistance to flow through the conduit, reduces the flow
volume, increases pump power consumption, decreasing the efficiency
of industrial operations. Further, this biomass can serve as a
source of contamination to flowing water or oil. Additionally, most
biofilms are heterogeneous in composition and structure which leads
to the formation of cathodic and anodic sites within the underlying
conduit metal thereby contributing to corrosion processes.
Currently, for pipeline treatment of biofilms, there is a trend to
use strong oxidizing biocides such as chlorine dioxide in cooling
systems and ozone in water distribution systems since low levels of
chlorine have been found to be ineffective against biofilms. Also,
a number of non-oxidizing biocides are available, which are
effective but their long-term effects on the environment are still
unclear. New techniques for biofilm control, such as ultrasound,
electrical fields, hydrolysis of EPS and methods altering biofilm
adhesion and cohesion are still in their infancy at the laboratory
level and are yet to be successfully demonstrated in large
industrial systems (Sriyutha Murthy P and Venkatesan R, "Industrial
Biofilms and Their Control". In: Marine and Industrial Biofouling;
Editors: Fleming H, Murthy P, Venkatesan R, and Cooksey K;
Springer-Verlag, 2010.).
One of the major economic losses faced by the oil and gas companies
is due to pipeline corrosion. The internal corrosion of the
pipelines is basically caused by sulfate reducing bacteria (SRB).
SRB are anaerobic and responsible for most instances of accelerated
corrosion damage. For biofilms created by SRB, some newer
strategies include the use of: a) calcium or sodium nitrates which
encourage more benign nitrate reducing bacteria to compete with
SRB, b) molybdate as a metabolic inhibitor preventing sulfate
reduction, c) anthraquinone which prohibits sulfide production and
its incorporation into the biofilm, and d) dispersants such as
filming amine technology which prevent biofilm adhesion. Also,
since there is no continuous water phase in oil pipelines (under
typical flow conditions) by which to dose bactericides, the use of
water-oil emulsions have been suggested ("Petroleum Microbiology";
edited by Ollivier B and Magot M, ASM Press, 2005.).
An example of the more recent biofilm altered adhesion concepts
includes the disclosure of International Patent Application
PCT/US2006/028353 describing a non-toxic, peptide-based biofilm
inhibitor that prevents Pseudomonas aeruginosa colonization of
stainless steel (and likely a wide variety of other metal surfaces)
and non-metalic surfaces. The compositions and methods describe a
very high affinity peptide ligand that binds specifically to
stainless steel and other surfaces to prevent Pseudomonas biofilm
formation. Another example of an inhibitor of biofilm adhesion is
the technology being developed by Australian firm BioSignal Ltd.
involving the use of furanones from the red seaweed Delisea
pulchra, which effectively avoids a broad spectrum of bacterial
infections without inciting any bacterial resistance to its
defensive chemistry. Furanones produced by this seaweed, bind
readily to the same specific protein-covered bacterial receptor
sites that receive the bacterial signaling molecules (N-acyl
homoserine lactone) which normally induce surface colonization.
BioSignal Ltd. is targeting the use of synthetic furanones to block
bacterial communication and thereby prevent bacteria from forming
groups and biofilms in applications including pipelines, HVAC, and
water lines treatment.
Methods of Decontamination of Food Processing, Storage, and
Transport Systems in the Food Industry
In addition to the more conventional means of decontamination
discussed above for other industrial applications, recently, the
food industry has embarked upon the use of enzyme-based schemes
that have been carried over from the bio-processing of food stuffs.
Specifically, efforts have been undertaken to find ways to
enzymatically degrade the EPS itself and thereby contribute to the
removal of biofilms. Largely, these efforts have been directed at
destruction of the polysaccharide framework of the EPS. A premier
example is found in the U. S. Patent Application 20110104141 to
Novozyme which discloses the use of alpha-amylase as a primary
enzyme for the breakdown of biofilm polysaccharides with the
potential inclusion of additional enzymes such as aminopeptidase,
amylase, carbohydrase, carboxypeptidase, catalase, cellulase,
chitinase, cutinase, cyclodextrin glycosyltransferase,
deoxyribonuclease, esterase, alpha-galactosidase,
beta-galactosidase, glucoamylase, alpha-glucosidase,
beta-glucosidase, haloperoxidase, invertase, laccase, lipase,
mannosidase, oxidoreductases, pectinolytic enzyme,
peptidoglutaminase, peroxidase, phytase, polyphenoloxidase,
proteolytic enzyme, ribonuclease, transglutaminase, or xylanase.
Products such as Biorem produced by Realco in coordination with
Novozyme to target applications in the food and beverage industry
exploit a two step cleaning process that invokes use of this kind
of multienzyme mixture followed by application of a biocide.
In this industrial sector also, ultrasound has been found a useful
tool; for sanitary control, it was found that the combination of
chelating agents with ultrasound has been useful for removing
selected biofilm-producing pathogens from metal surfaces (Oulahal
N, Martial-Gros A, Bonneau M and Blum L J, "Combined effect of
chelating agents and ultrasound on biofilm removal from stainless
steel surfaces. Application to "Escherichia coli milk" and
"Staphylococcus aureus milk" biofilms", Biofilms, 2004; 1: 65-73,
Cambridge University Press.). The efficacy of such ensonification
has been shown to exhibit dependency on the frequency and duty
cycle of the energy (Nishikawa T, et al., "A study of the efficacy
of ultrasonic waves in removing biofilms," Gerontology, September
2010; Vol 27, Issue 3: 199-206.).
Current Methods for Treating Marine Biofouling
Biofouling occurs worldwide in various industries and one of the
most common biofouling sites is on the hulls of ships, where
barnacles are often found. A significant problem associated with
biofilms on ships is the eventual corrosion of the hull, leading to
the ship's deterioration. However, before corrosion occurs, organic
growth can increase the roughness of the hull, which will decrease
the vessel's maneuverability and increase hydrodynamic drag.
Ultimately, biofouling can increase a ship's fuel consumption by as
much as 30%. Parts of a ship other than the hull are affected as
well: heat exchangers, water-cooling pipes, propellers, even the
ballast water. Fishing and fish farming are also affected, with
mesh cages and trawls harboring fouling organisms. In Australia,
biofouling accounts for about 80% of the pearling industry's costs
(Stanczak M, "Biofouling: It's Not Just Barnacles Anymore," CSA
Discovery Guide, 2004;
http://www.csa.com/discoveryguides/biofoul/overview.php.).
The traditional method of control is to coat exposed surfaces with
an anti-fouling compounds. Most of these compounds rely on copper
and tin salts that gradually leach from the coating and contaminate
the surrounding environment. One of the most widely used coatings
to date has been tributyl tin (TBT) which is highly toxic to marine
organisms. Since it has been found to have unwanted side-effects on
non-target organisms, a world-wide ban on its use was instituted in
2008. The race is on for an environmentally sound alternative
(Scottish Association for Marine Science,
http://www.sams.ac.uk/researchldepartments/microbial-molecular/m-
mb-project-themes/algal-biofilms.).
Hence, in maritime applications such as shipping, there is an unmet
need for viable, cost-effective biofilm remediation.
Current Methods for Treating Biofilms in Heating, Ventilation and
Air-Conditioning (HVAC) and Refrigeration Systems
HVAC and refrigeration systems encounter problems associated with
biofilms formed on cooling coils, drain pans, and in duct work
subjected to water condensation. Biofilm formation on cooling coils
diminishes heat exchange efficiency; its growth on other surfaces,
including drain pans and duct work, is a source of contamination in
the air stream. Conventional methods of addressing biofilms in
these applications include maintenance cleaning of coils, duct work
and drain pans, use of anticorrosion and antimicrobial coatings on
system surfaces, and the exposure of system surfaces to C-band
ultraviolet light to break down biofilms and kill pathogens.
Remediation of Biofilm Contamination in Household Applications
The household products industry is vitally concerned with
disinfection of household surfaces, water and plumbing systems, and
human hygienic needs. Difficulties associated with killing bacteria
attached to these diverse surfaces are well known in this
industrial sector and considerable research currently is directed
at developing products which kill or remove biofilms.
An innovation in this sector is probiotic-based cleaning. Some
versions of these products lay down layers of benign bacteria that
successfully compete with pathogenic bacteria for resources on
kitchen and bathroom surfaces. Other such products combine enzymes
with probiotic bacteria to digest biofilms and dead pathogens. A
leading example of this class of products is PIP produced by
Chrisal Probiotics of the Netherlands.
The conventional approaches to treatment of biofilm discussed for
both medical and industrial applications variously have been
unproven, of limited effectiveness, time consuming, costly in cases
where large surface areas are involved or surfaces require repeated
treatment, and newer concepts have yet to demonstrate effectiveness
and scalability to field applications. Hence, there remains an
urgent need for more effective and less costly methods to treat
biofilms. The present compositions and method offer the prospect of
a new standalone approach to biofilm treatment with higher efficacy
and lower cost, with additional potential for augmenting certain
conventional treatments while reducing the costs of such
treatments.
SUMMARY
Trehalose (a universal general stress response metabolite and an
osmoprotectant) can play an important role in the formation and
development of microbial biofilm and the specific interactions of
trehalose with water can be considered to be one of the most
important mechanisms of biofilm formation. The present compositions
and methods have been conceived to target trehalose degradation as
a key step in degrading biofilm.
In various embodiments of the compositions and methods, compounds
that prevent, degrade, and/or inhibit the formation of biofilms,
compositions comprising these compounds, devices exploiting these
compounds, and methods of using the same are disclosed.
Because trehalose serves to manipulate hydrogen bonds among water
molecules and bacterial cells in the process of forming the biofilm
gel, the degradation of trehalose ultimately should result in
degradation of the biofilm gel. A class of compounds that degrade
trehalose with high specificity, thereby degrading the biofilm
matrix gel is disclosed. Specifically, the naturally occurring
enzyme trehalase will hydrolyze a molecule of trehalose into two
molecules of glucose. The small amount of enzyme trehalase produced
in the human body must be augmented with the administration of much
larger amounts to treat in vivo biofilm-based infections. Various
treatment formulations that incorporate trehalase enzymes and
associated delivery mechanisms are detailed for specific types of
infections; these include systemic and local treatment protocols.
Additionally, trehalase-containing mixtures and associated
processes are disclosed to degrade biofilms present on medical
instruments and to mitigate biofilm fouling and biofilm-based
biocorrosion for industrial applications. For degrading biofilms on
medical equipment, trehalase-containing mixtures can be used in
concert with other processes, such as ultrasound and
ultrasound-assisted enzymatic activity to degrade biofilms. Biofilm
prevention approaches comprise the use of trehalase enzymes in
surface coatings.
Following is a lexicon of terms and phrases that more particularly
define the compositions and methods and support the meaning of the
claims:
Time-delayed release--in the context of the present compositions
and methods, time-delayed release refers specifically to trehalase
(or other compounds) release that occurs at a predetermined
approximate time after the trehalase (and in some embodiments,
other compounds) in pill, capsule, tablet or other form is ingested
orally. Typically, for the present compositions and methods, the
time delay means that the initial release of trehalase (or other
compounds) will occur in the small intestine, to avoid degradation
by naturally occurring proteolytic enzymes in the upper GI tract.
Various pre-programmed temporal profiles for release in the small
intestine are within the scope of the compositions and methods,
such as, for example, linearly increasing or decreasing rates of
release with time, or a constant rate of release.
Sustained release--in the context of the present compositions and
methods, it refers to the release of trehalase (or other compounds)
for applications external to the body. This is a continuous release
of trehalase (or other compounds) that is not time-delayed, but is
initiated at first opportunity for the purpose of continuous,
ongoing exposure of medical device and industrial surfaces to
treatment enzymes.
Sufficient for efficacy--pertains to treatment composition amounts
and treatment exposure durations adequate to breakdown the gel
structure of biofilm for its dispersal and further penetration by
antimicrobial agents to treat the target infectious pathogens.
Trehalase--refers to any enzyme selected from the category of
trehalase isoenzymes. There are two types of trehalase enzymes
found in microorganisms: neutral trehalase (NT) typically found in
the cytosol and acid trehalase (AT) found in the vacuoles of the
cytosol, either of which type may find application in the present
compositions and methods. Further, the number of candidate enzymes
is large; as many as 541 model variants (isoenzymes) of trehalase
can be found in the Protein Model Portal
(http://www.proteinmodelportal.org), each exhibiting varying
potencies in the hydrolysis of trehalose into glucose. The present
compositions and methods anticipate a selection from among these
isoenzymes that is optimized for the specific biofilm application.
For example, the ability to sufficiently purify a given isoenzyme
for internal bodily use may favor its selection for this purpose
over another isoenzyme that exhibits higher enzymatic activity, but
which would be relegated to industrial applications.
Digestive enzymes--are enzymes that break down polymeric
macromolecules of ingested food into their smaller building blocks,
in order to facilitate their absorption by the body. In the present
icompositions and methods, treatment formulations comprising
trehalase (or other compounds) are disclosed which should: a) avoid
degradation by the digestive enzymes naturally occurring in the
upper GI tract and b) be combined in time-delayed release form with
digestive enzyme supplements to avoid degradation by proteolytic
enzymes in such supplements.
Medical devices--comprise devices that are installed either
temporarily or permanently in the body and medical instruments that
may or may not contact the body, but at least contact tissue or
bodily fluids. Examples of temporarily installed medical devices
include catheters, endoscopes, and surgical devices. Permanent
devices examples include devices such as orthopedic implants,
stents, and surgical mesh. Examples of devices used external to the
body include stethoscopes, dialysis machines, and blood and urinary
analysis instruments. Each of the aforementioned devices exhibit
surfaces that are vulnerable to biofilm formation and therefore can
benefit from treatment by specific embodiments of the presently
disclosed compositions and methods.
Antimicrobials--are substances that kill or inhibit the growth of
microorganisms such as bacteria, fungi, or protozoans.
Antimicrobials either kill microbes (microbiocidal) or prevent the
growth of microbes (microbiostatic). Disinfectants are
antimicrobial substances used on non-living objects or outside the
body.
Other saccharidases (enzymes hydrolyzing saccharides)--include
various di-, oligo-, and polysaccharidases.
Living organisms--pertains to the spectrum of living entities from
microbes to animals and humans.
GI tract--refers to the gastrointestinal tract; the upper GI tract
comprising the mouth, esophagus, stomach, and duodenum, and the
lower GI tract comprising the small and large intestines.
Administering via the GI tract--relates to three main alternative
treatment delivery methods: first is oral administration in which
the treatment compounds are administered via the mouth; for the
patients that may not be able to receive treatment by mouth, the
second method available is by the naso-gastric tube; and a third
method includes delivery by colonic irrigation.
Administering via systemic use--relates to administration of
treatment compounds by percutaneous injection, intramuscular
injection, intra-venous injection, and venous catheter
administration.
Other aspects, advantages, and features of the present disclosure
will become apparent after review of the entire application,
including the following sections: Brief Description of the
Drawings, Detailed Description, and the Claims.
BRIEF DESCRIPTION OF THE DRAWINGS
FIG. 1A is diagram of the chemical structure of the dissacharide
trehalase.
FIG. 1B is a pictorial diagram of the backbone structure of
trehalase.
FIG. 2A is a ribbon model pictorial diagram of an enzyme of
trehalase derived from Saccharomyces cerevisiae.
FIG. 2B is a ribbon model pictorial diagram of an enzyme of
trehalase derived from Penicillium marneffei.
FIG. 2C is a ribbon model pictorial diagram of an enzyme of
trehalase derived from Homo sapiens.
FIG. 2D is a ribbon model pictorial diagram of an enzyme of
trehalase derived from Candida albicans.
FIG. 3 is a summary chart showing the biofilm produced by P.
aeruginosa PAO1 in accordance with a non-limiting example.
FIG. 4 is a summary chart of the biofilm produced by S. aureus
ATCC25923 in accordance with a non-limiting example.
FIG. 5 is another summary chart of the biofilm produced by S.
aureus ATCC25923 in accordance with a non-limiting example.
FIG. 6 is another summary chart of the biofilm produced by P.
aeruginosa PAO1 in accordance with a non-limiting example.
FIG. 7 is a summary chart for the results of the MIC determination
with selected S. Aureus strains in accordance with a non-limiting
example.
FIG. 8 is a summary table for the results of the screening of
clinical isolates in accordance with a non-limiting example.
FIGS. 9A through 9C are charts for the results of the Trehalase
testing on clinical isolates where the biomass has crystal violet
staining in accordance with a non-limiting example.
FIGS. 10A through 10C are charts for the results of the Trehalase
testing on clinical isolates for cell viability with resazurin
staining in accordance with a non-limiting example.
FIGS. 11A through 11C are charts for the results showing the
effectiveness of Trehalase added during biofilm growth on a
catheter segment up to 24 hours in accordance with a non-limiting
example.
FIGS. 12A through 12C are charts for the results of the Trehalase
added after 24 hours biofilm growth on a catheter segment in
accordance with a non-limiting example.
DETAILED DESCRIPTION
Since any bacterial biofilm can be defined as a living dynamic
structure with spatial and temporal heterogeneity for both
components of microbial biofilm (the exopolymer gel matrix and the
microcolonies of infectious microorganisms embedded in this gel
matrix), the treatment approaches for biofilm-based chronic
infections should be aimed simultaneously at both components of
microbial biofilm: prevention of formation, inhibition of growth
and degradation of biofilm gel matrix, and killing the infectious
pathogens embedded in the biofilm gel matrix. In this context,
trehalase should be included into existing approaches for
prevention and treatment of microbial biofilms, being used in
combination with existing natural and synthetic amtimicrobials and
anti-biofilm substances and methods of their use to increase their
effectiveness. A review of recent patent literature summarizes
citations under six categories of current treatment approaches: 1)
antibiotics and small molecule inhibitors of new and established
biofilms and biofilm-forming infectious pathogens, 2) quorum
sensing and signaling molecules inhibitors, 3) surface coating
substances for inhibition of biofilm formation on medical devices
and equipment, 4) antibodies and vaccines for infectious biofilm
treatment, 5) enzymes for degrading biofilms, and 6) bacteriophage
treatment of infectious biofilms (Lynch A S and Abbanat D, "New
antibiotic agents and approaches to treat biofilm-associated
infections," Expert Opin. Ther. Patents, 2010; 20(10):
1373-1387.).
All microorganisms in their natural environments encounter a
multitude of stresses, including osmotic stress, oxidative stress,
membrane and cell envelope stress, ribosomal stress, nutrient and
oxygen limitations, temperature stress, and many other stresses as
characteristics of various specific environmental conditions.
Infectious pathogens encounter the same stresses in their
environment--the internal milieu of their host (including human
body). The infectious process of biofilm-based infections in the
human body should be considered as a constant battle between
infectious microorganism (for its adaptation, survival and
proliferation in the milieu of a human body) and a host who uses
all its natural defense mechanisms to kill the pathogen and
eliminate it from the body, restore normal homeostasis, and repair
damaged body tissues. The exposure to various stresses, impact
bacterial susceptibility to a variety of antimicrobials through
their initiation of stress responses that recruit various
resistance determinants or promote physiological changes that
compromise antimicrobial activity (Keith Poole, "Bacterial stress
responses as determinants of antimicrobial resistance", J
Antimicrob Chemother 2012; 67: 2069-2089.). One of the most
important survival mechanisms for any bacteria in any environment
is the general stress response (ubiquitous in nature) triggered by
various environmental stresses and their interactivities via
genetic regulation. In general stress response, the increased
production of trehalose (as a general stress response metabolite
and an osmoprotectant) plays an important role in adaptation and
survival of infectious pathogen in the host body, and plays an
important role in antimicrobial resistance as well.
Trehalose is a disaccharide that is ubiquitous in the biosphere and
present in almost all forms of life except mammals. In various
bacteria and fungi, it is one of the most important storage
carbohydrates, serving as a source of energy and as a carbon source
for synthesis of cellular components; it can play a transport role
and control certain metabolic pathways; it functions as a
protectant for cell membranes and cell proteins against the adverse
effects of various stresses, including the osmotic stress, heat,
cold, desiccation, dehydration, deprivation of nutrients,
oxidation, and anoxia (Elbein A D, "The metabolism of
.alpha.,.alpha.-trehalose," Adv. Carbohyd. Chem. Biochem., 1974;
30: 227-256.); (Crowe J, Crowe L, and Chapman D, "Preservation of
membranes in anhydrobiotic organisms. The role of trehalose,"
Science, 1984; 223: 209-217.); (Takayama K and Armstrong E L,
"Isolation, characterization and function of
6-mycolyl-6'acetyltrehalose in the H37Rv strain of Mycobacterium
tuberculosis," Biochemistry, 1976; 15: 441-446.); (Christopher
Askew, Adnane Sellam, Ellias Epp, Herve Hogues, Alaka Mullick,
Andre Nantel, Malcolm Whiteway, "Transcriptional regulation of
carbohydrate metabolism in the human pathogen Candida albicans",
PLoS Pathogens 2009-10-01.); (Joke Serneels, Helene Tournu, Patrick
Van Dijck, "Tight control of trehalose content is required for
efficient heat-induced cell elongation in Candida albicans",
Journal of Biological Chemistry, 2012-10-26.).
Trehalose may be partially responsible for the virulence and
antimicrobial resistance properties in various opportunistic and
pathogenic microorganisms, including those known to cause chronic
infections with biofilm formation in the human body, including:
Pseudomonas spp., Bacillus spp., Staphylococci spp., Streptococci
spp, Haemophilus influenza, Klebsiella pneumoniae, Proteus spp.,
Mycobacteria spp., Corynebacteria spp., Enterococci spp.,
enteropathogenic E. coli, various human pathogenic yeasts and fungi
(Candida spp., Cryptococcus neoformans, Aspergillus spp.). As
demonstrated in some strains of Candida albicans, interference with
the production of trehalose strongly reduces their virulence.
Specifically, C. albicans mutants with deleted gene TSP2, which
encodes trehalose-6-phosphate phosphatase, one of two enzymes
involved in trehalose synthesis, exhibited diminished virulence in
in vivo mouse model of systemic infection and, being grown within
in vitro biofilm systems, displayed significantly less biofilm
formation than selected non-mutant strains (Coeney T, Nailis H,
Tournu H, Van Dick P, and Nelis H, "Biofilm Formation and Stress
Response in Candida Albicans TSP2 Mutant," ASM Conference on
Candida and Candidiasis, Edition 8, Denver, Colo.; Mar. 12-17,
2006.). The in vivo studies in the pathobiology of Cryptococcus
neoformans, identified the presence of a functioning trehalose
pathway during experimental infection in the mouse and rabbit
models and suggested its importance for C. neoformans survival in
the host; using created null-mutants of the trehalose-6-phosphate
(T6P) synthase (TPS1), trehalose-6-phosphate phosphatase (TPS2),
and neutral trehalase (NTHJ) genes, it was demonstrated that both
TPS1 and TPS2 are required for high-temperature (37.degree. C.)
growth and glycolysis, but the block at TPS2 results in the
apparent toxic accumulation of T6P, which makes the enzyme
trehalose-6-phosphate phosphatase a fungicidal target (Elizabeth
Wills Petzold, Uwe Himmelreich, Eleftherios Mylonakis, Thomas Rude,
Dena Toffaletti, Gary M. Cox, Jackie L. Miller, and John R.
Perfect, "Characterization and Regulation of the Trehalose
Synthesis Pathway and Its Importance in the Pathogenicity of
Cryptococcus neoformans", Infection and Immunity, October 2006, p.
5877-5887.).
All microorganisms can synthesize trehalose intracellularly and/or
take it from the environment using various synthesis and
degradation pathways for trehalose metabolism. The specific use of
these pathways by various microorganisms depends on their genetic
program for trehalose utilization and availability of substrates
for trehalose biosynthesis in their environment.
Many microorganisms, including human pathogenic bacteria and fungi,
synthesize trehalose intracellularly mostly via pathways that
utilize various nucleoside diphosphate glucose derivatives as
glucosyl donors (ADP-D-glucose, CDP-D-glucose, GDP-D-glucose,
TDP-D-glucose and UDP-D-glucose) and a-D-glucose-6-phosphate in a
two-step reaction: in the first step--formation of intermediate
metabolite trehalose 6-phosphate (T6P) by the action of enzyme
trehalose-6-phosphate synthase (TPS), and in the second
step--formation of final product trehalose (a, a-trehalose), by the
action of enzyme trehalose-6-phosphate phosphatase (TPP) (Styrvold
O B and Strom A R, "Synthesis, accumulation, and excretion of
trehalose in osmotically stressed Escherichia coli K-12 strains:
influence of amber suppressors and function of the periplasmic
trehalase," J. Bacteriol, 1991; 173(3): 1187-1192. PMID: 1825082).
It should be mentioned that some mycobacteria, such as
Mycobacterium smegmatis and Mycobacterium tuberculosis, possessing
unusual trehalose-6-phosphate synthases, are capable of utilizing
all five nucleoside diphosphate glucose derivatives as glucosyl
donors (Lapp D, Patterson B W, Elbein A D, "Properties of a
trehalose phosphate synthetase from Mycobacterium smegmatis.
Activation of the enzyme by polynucleotides and other polyanions,"
J. Biol. Chem., 1971; 246 (14): 4567-4579.).
Also, many microrganisms, can synthesize trehalose directly from
disaccharide maltose (degradation product of glycogen and starch)
independently of the presence of phosphate compounds
trehalose-6-phosphate and glucose-6-phosphate. This pathway
involves the intramolecular rearrangement of maltose
(glucosyl-alpha1,4-glucopyranoside) to convert the 1,4-linkage into
the 1,1-linkage of trehalose by the action of enzyme trehalose
synthase (TS), forming free trehalose (a, a-trehalose) as the
initial product. It is postulated that in Corynebacterium
glutamicum this pathway may work in the opposite direction,
compensating for the absence of a trehalase enzyme, by converting
excess trehalose back into maltose, for reuse as a carbon source
(De Smet K A, Weston A, Brown I N, Young D B, Robertson B D, "Three
pathways for trehalose biosynthesis in mycobacteria," Microbiology,
2000; 146 (Pt 1): 199-208. PMID: 10658666); (Wolf A, Cramer R,
Morbach S, "Three pathways for trehalose metabolism in
Corynebacterium glutamicum ATCC 13032 and their significance in
response to osmotic stress," Mol Microbiol, 2003; 49(4): 1119-1134.
PMID: 12890033.).
In an additional pathway, trehalose can be formed from
polysaccharides (such as glycogen or starch) in multi-step process
by the action of several enzymes: in the first step--the enzyme
isoamylase hydrolyzes the .alpha.-1,6-glucosidic linkage in
glycogen or the a-1,4-glucosidic linkages in other polysaccharides
(such as starch from plants), to produce a maltodextrin
(oligosaccharide); in the next step--the enzyme
maltoolgosyl-trehalose synthase (MTS) converts maltodextrin to
maltooligosyl-trehalose by forming an
.alpha.,.alpha.-1,1-glucosidic linkage via intermolecular
transglucosylation; and in the third step--the enzyme
maltooligosyl-trehalose trehalohydrolase (MTTH) hydrolyzes the
product, forming free trehalose (a, a-trehalose) and a maltodextrin
which becomes shorter by two glucosyl residues. In Corynebacterium
glutamicum, which possess three different pathways for trehalose
biosynthesis, this is the main route for trehalose biosynthesis
(Maruta K, Mitsuzumi H, Nakada T, Kubota M, Chaen H, Fukuda S,
Sugumoto T, Kurimoto M, "Cloning and sequencing of a cluster of
genes encoding novel enzymes of trehalose biosynthesis from
thermophilic archaebacterium Sulfolobus acidocaldarius," Biochim
Biophys Acta, 1996; 129 (3): 177-181. PMID: 8980629.).
For degradation of trehalose, various microorganisms, including
human pathogenic bacteria and fungi, can utilize several
alternative pathways. Unmodified trehalose (a, a-trehalose) may be
degraded by a hydrolyzing enzyme trehalase (a, a-trehalohydrolase),
yielding two .beta.-D-glucose molecules, or it may be split by the
action of the enzyme trehalose phosphorylase (TP), yielding
.beta.-D-glucose-6-phosphate as the end product. Trehalose
phosphorylase (TP), can also catalyze the reversible synthesis and
degradation of trehalose from/to a .beta.-D-glucose-1-phosphate and
.beta.-D-glucose, or .alpha.-D-glucose-1-phosphate and
.alpha.-D-glucose. Phosphorylated form, trehalose-6-phosphate, can
be either hydrolyzed by trehalose-6-phosphate hydrolase, yielding
.beta.-D-glucose and .beta.-D-glucose-6-phosphate, or degraded by
trehalose-6-phosphate phosphorylase, yielding
.beta.-D-glucose-1-phosphate and .beta.-D-glucose-6-phosphate. All
end products of the degradation pathways can be metabolized via
glycolysis. All end products of trehalose degradation pathways can
be metabolized via glycolysis. (Helfert C, Gotsche S, Dahl M K,
"Cleavage of trehalose-phosphate in Bacillus subtilis is catalyzed
by a phospho-alpha-(1-1)-glucosidase encoded by the TreA gene," Mol
Microbiol, 1995; 16(1): 111-120. PMID: 7651129.); (Levander F,
Andersson U, Radstrom P, "Physiological role of
beta-phosphoglucomutase in Lactococcus lactis," Appl Environ
Microbiol, 2001; 67(10): 4546-4553. PMID: 11571154.).
For survival in the live environment of a human body, the
pathogenic microorganisms must continuously adapt to temporal and
spatial fluctuations in osmolarity of body fluids. In
osmoadaptation, bacteria constitutively use the universal mechanism
of uptake and release of osmotically active compounds (osmolytes).
Bacteria adapt to the conditions of increased external osmolarity
by importing charged ions from the environment, and importing or
synthesizing compatible solutes. Upon a shift to a low-osmolarity
media, the excretion of osmolytes is required to restore normal
turgor and prevent the cells from bursting. The pathways for import
and efflux of compatible solutes include PTS system, ABC
transporters, mechanosensitive channels, and porins (Berrier C M,
Besnard M, Ajouz B, Coulombe A, and Ghazi A, "Multiple
mechanosensitive ion channels from Escherichia coli, activated at
different thresholds of applied pressure," J. Membr. Biol., 1996;
151: 175-187.); (Bremer R and Kraemer R, "Coping with osmotic
challenges: osmoregulation through accumulation and release of
compatible solutes in bacteria," pp. 79-97. In G. Storz and R.
Hengge-Aronis (ed.), Bacterial stress responses. 2000; ASM Press,
Washington, D.C.); (Chang G, Spencer R, Lee A T, Barclay M T, and
Rees D C, "Structure of the MscL homolog from Mycobacterium
tuberculosis: a gated mechanosensitive ion channel," Science, 1998;
282: 2220-2225.); (Morbach S and Kraemer R, "Body shaping under
water stress: osmosensing and osmoregulation of solute transport in
bacteria," ChemBioChem, 2002; 3: 384-397.).
Compatible solutes are small, zwitterionic, highly soluble organic
molecules, which include diverse substances, such as amino acids
(proline,glutamate), amino acid derivatives (glycine betaine,
ectoine), and sugars (trehalose and sucrose), that are thought to
stabilize proteins and lead to the hydration of the cell (Steator R
D and Hill C, "Bacterial osmoadaptation: the role of osmolytes in
bacterial stress and virulence," FEMS Mocrobiol. Rev., 2002; 26:
49-71.). Various bacteria may prefer different osmolytes taken from
the environment, but all of them constitutively utilize trehalose
(taken from the environment or synthesized intracellularly) as a
universal osmoprotectant. For example, E. coli and Vibrio Cholerae
in human GI tract prefer glycine betaine, but its synthesis relies
on an external supply of proline, betaines, or choline which may
not be readily available in the environment or significantly
reduced in the deeper layers of microbial biofilm. When these
compounds are not available, a microbial cell can achieve a
moderate level of osmotic tolerance by accumulation of glutamate
and trehalose (Styrvold O B, Strom A R, "Synthesis, accumulation,
and excretion of trehalose in osmotically stressed Escherichia coli
K-12 strains: influence of amber suppressors and function of
periplasmic trehalase," J Bacteriol, 1991; 173 (3): 1187-1192.
PMID: 1825082.); (Kapfhammer D, Karatan E, Pflughoeft K J, and
Watnik P I, "Role for Glycine Betaine Transport in Vibrio cholera
Osmoadaptation and Biofilm Formation within Microbial Communities,"
Applied and Environmental Microbiology, July 2005: 3840-3847.).
As demonstrated in laboratory-grown bacteria, the first adaptive
response to osmotic stress comprises both the increased uptake rate
and the amount of cytosolic potassium, followed by the accumulation
of glutamate and synthesis of trehalose (Dinnbier U, Limpinsel E,
Schmid R, and Bakker E P, "Transient accumulation of potassium
glutamate and its replacement by trehalose during adaptation of
growing cells of Escherichia coli K-12 to elevated sodium chloride
concentrations," Arch. Microbiol., 1988; 150: 348-357.); (McLaggan
D, Naprstek J, Buurman E T, and Epstein W, "Interdependence of
K.sup.+ and glutamate accumulation during osmotic adaptation of
Escherichia coli," J. Biol. Chem., 1994; 269: 1911-1917.); (StromAR
and Kaasen I, "Trehalose metabolism in Escherichia coli: stress
protection and stress regulation of gene expression," Mol.
Microbiol., 1993; 8: 205-210.). The time-dependent (10 to 60
minutes) alterations in the proteome of E. coli (grown under
aerobic conditions) in response to osmotic stress, demonstrated
upregulated genes for synthesis of both trehalose and cytosolic
trehalase (trehalose-degrading enzyme with regulatory properties)
in the middle phase (10 to 30 minutes) and in the long phase (30 to
60 minutes) of bacterial adaptation to hyperosmotic stress, with
the trehalase synthesis genes (TreF) upregulated in the early phase
of adaptation (0 to 10 minutes) (Weber A, Kogl S A, and Jung K,
"Time-Dependent Proteome Alterations under Osmotic Stress during
Aerobic and Anaerobic Growth in Escherichia coli," Journal of
Bacteriology, October 2006: 7165-7175. doi:
10.1128/JB.00508-06.).
Trehalose is a stable dissacharide with glycosidic bond
[O-.alpha.-D-Glucopyranosyl-(1-1)-.alpha.-D-glucopyranoside] formed
from a condensation between the hydroxyl groups of the anomeric
carbons of two molecules of glucose, preventing them from
interacting with other molecules and thereby rendering trehalose
among the most chemically inert sugars (Birch G G, "Trehaloses,"
Adv. Carbohydr. Chem. Biochem., 1963; 18: 201-225.); (Elbein A D,
"The metabolism of alpha, alpha-trehalose," Adv. Carbohydr. Chem.
Biochem., 1974; 30: 227-256.). The flexible glycosidic bond,
together with the absence of internal hydrogen bonds, yields a
supple molecule, but this glycosidic bond does not break easily:
the lkcal/mol linkage is highly resilient, enabling the trehalose
molecule to withstand a wide range of temperature and pH conditions
(Pava C L and Panek A D, "Biotechnological applications of the
disaccharide trehalose," Biotechnol. Annu. Rev., 1996; 2:
293-314.). Because of the unusual glycosidic bond between the
anomeric carbons (1-1), there are no more accessible carbons for
further polymerization, so that trehalose exists only as a
disaccharide, being rather distributed as disaccharide molecules in
the gel-like matrix of biofilm, influencing its density via
interaction between trehalose and water molecules. Intermolecular
hydrogen bonds (H bonds), the strongest of intermolecular forces,
are central to trehalose interaction with water. Specifically, such
bonds modify the structure of water surrounding trehalose molecules
and account for the self-aggregation phenomena of trehalose
molecules observed in molecular dynamic simulations and supported
by experimental studies.
The chemical structure of trehalose is depicted in FIG. 1a
indicating an alpha-linked disaccharide formed by an
.alpha.,.alpha.-1,1-glucosidic bond between two .alpha.-glucose
units. The backbone structure of this enzyme is shown in FIG. 1b
depicting the two planes established by the glucose units.
Trehalose has a unique ability to capture water through extensive
solvation. Water molecules are arranged in a solvation complex
around trehalose molecules, with water associating with trehalose
functional groups through H bond formation; at infinite dilution,
the solvation number approaches 15 (the highest among all
disaccharides). Trehalose is able to restructure water even at
minimum aqueous concentrations, supporting the gelation phenomena
in these conditions. With respect to water restructuring behavior,
trehalose enhances the hydrogen bonding between water molecules by
approximately 2%. This is sufficient to destructure the pure water
tetrahedral network in conformity with a restructuring imposed by
trehalose clusters. Stronger, more linear, and better optimized H
bonds are formed between water molecules, while weaker bonds are
relegated to trehalose-water interactions (Sapir L and Harries D,
"Linking Trehalose Self-Association with Binary Aqueous Solution
Equation of State," J. Phys. Chem. B, 2011; 115: 624-634.).
Trehalose self-associates in aqueous solutions in a concentration
dependent manner to form clusters of increasing size, until finally
forming percolating, infinitely connected, clustering networks (at
concentrations of 1.75 M and higher), affecting the dynamic
properties of the solution. The lack of intramolecular hydrogen
bonds in trehalose, compared with other disaccharides (sucrose,
maltose, isomaltose), accounts for its higher tendency to
aggregate, thereby already affecting the dynamic properties of
water at lower trehalose concentrations (Lebret A, Bordat P,
Affouard F, Descamps M, Migliardo F J, Phys. Chem. B, 2005; 109:
11046.); (Lebret A, Affouard F, Bordat P, Hedoux A, Guinet Y, and
Descamps M, Chem. Phys., 2008; 345:267.); (Peric-Hassler L, Hansen
H S, Baron R, and Hunenberger P H, "Conformational properties of
glucose-based disaccharides investigated using molecular dynamics
simulations with local elevation umbrella sampling," Carbohydr.
Res., 2010; 345: 1781.)
In a ternary mixture of protein (lysozyme), sugar, and water, at a
moderate concentration of 0.5 M, trehalose can cluster around the
protein, thereby trapping a thin layer of water molecules with
modified solvation properties, playing the role of a "dynamic
reducer" for solvent water molecules in the hydration shell around
the protein. A remarkable conformational rigidity of the trehalose
molecule due to anisotropic hydration (very little hydration
adjacent to the glycosidic oxygen of trehalose), provides stable
interactions with hydrogen-bonded water molecules; trehalose makes
an average of 2.8 long-lived hydrogen bonds per each step of
molecular dynamic simulation compared with the average of 2.1 for
the other sugars (Lins R D, Pereira C S, and Hunenberger P H,
"Protein-Trehalose Interactions in Aqueous Solution," Proteins,
2004; 55: 177.); (Choi Y, Cho K W, Jeong K, and Jung S, "Molecular
dynamic simulations of trehalose as a `dynamic reducer` for solvent
water molecules in the hydration shell," Carbohydr Res., Jun. 12,
2006; 341(8): 1020-1028.).
In a simulated ternary mixture of lipid membranes, composed of DPPC
(dipalmitoylphosphatidylcholine), with aqueous solution of
trehalose, the trehalose molecules cluster near membrane
interfaces, forming hydrogen bonds both between trehalose molecules
and with the lipid headgroups (Pereira C S, Hunenberger P H, "The
effect of trehalose on a phospholipid membrane under mechanical
stress," Biophys. J., 2008; 95: 3525.); (Sum A K, Faller R, and de
Pablo J J, "Molecular simulation study of phospholipid bilayers and
insights of the interactions with disaccharides," Biophys. J.,
2003; 85: 2830.). Trehalose may compete with water binding to both
carbonyls and phosphates in cell membranes, forming the OH bridges
that are stronger than the H-bonds of water with those groups, and
the displacement of water is compensated with the insertion of
sugar. Trehalose, a dimer of glucose with the ability to form at
least 10 hydrogen bonds, inserts in a lipid interface nearly normal
to the lipid bilayer plane and can decrease water activity in the
cell membrane up to 70% at a concentration of trehalose as low as
0.1 mM. The insertion of trehalose, replacing water simultaneously
at the carbonyls and the phosphates, does not cause the surface
defects in the cell membrane with respect to hydrated lipids
(Pereira C S and Hunenberger P H, "The effect of trehalose on a
phospholipid membrane under mechanical stress," Biophys. J., 2008;
95:3525.); (Sum A K, Faller R, and de Pablo J J, "Molecular
simulation study of phospholipid bilayers and insights of the
interactions with disaccharides," Biophys. J., 2003; 85: 2830.);
(Villareal M, Diaz S B, Disalvo E A, Montich G, Langmuir, 2004; 20:
7844.). We hypothesize that disaccharide trehalose, being inserted
into the phospholipid bilayer(s) of bacterial cell membrane(s) for
protection of their integrity, can affect the fluidity of the cell
membranes (via its specific interactions with water, carbonyls and
phosphates), and cause conformational changes in trans-membrane
proteins (including ion channels, efflux pumps and porins),
resulting in changes of their functional properties.
As a result of water displacement, trehalose may affect the cell
surface potential and hence microbial cell aggregation and
attachment to surfaces. There can be at least two mechanisms for
these phenomena. First, the magnitude of cell surface potential can
be modulated by trehalose displacement of water in its attachment
to cell membrane phospholipids and carbonyl compounds. Second, this
same displacement of water (in a non-unifonnrm manner) can lead to
heterogeneity of surface potential, also imparting the adhesion
properties of microorganism. (Poortinga A T, Bos R, Norde W, and
Busscher H J, "Electric double layer interactions in bacterial
adhesion to surfaces," Surface Science Reports, 2002; 47: 1-31.);
(Disalvo E A, Lairion F, Martini F, Almaleck H, Diaz S, and
Gordillo G, "Water in Biological Membranes at Interfaces: Does it
Play a Functional Role?," An. Asoc. Quim. Argent., 2004; V. 92 n.
4-6 Buenos Aires ago./dic.).
Trehalose and Biofilm Formation
Based on the unique properties of trehalose as a universal general
stress response metabolite and an osmoprotectant, and the specific
features of its interactions with water (which comprises up to 95%
of biofilm matrix), trehalose can be one of the most important
components of microbial biofilm, and its specific interactions with
water can be considered to be one of the most important mechanisms
of biofilm formation.
Since the formation of microbial biofilm can be seen as a
continuous process of adaptation of a microorganism to its
environment, trehalose and its interactions with water can play an
important role in all stages of microbial biofilm development (the
early stage of initial biofilm formation, maturation of the
biofilm, and dispersal of the biofilm).
From the first moment, when a microorganism enters the human body
in a planktonic form, it is subjected to various stresses (first of
all, osmotic stress) and undergoes the general stress response with
the production of trehalose, which in its initial interactions with
water begins the process of microbial biofilm formation. In this
initial stage, trehalose facilitates adhesion of planktonic
bacteria to surfaces by various means: as a result of its
interaction with water and the lipid headgroups at the cell
membrane interfaces, it decreases the microbial cell surface
potential and enhances the bacterial cell aggregation, initial
adsorption and attachment to the surfaces, both biotic and abiotic.
Also, trehalose favors the bacterial cell aggregation and
attachment to various surfaces by forming a hydration layer with
modified solvation properties around the bacterial cell and
reducing the dynamic properties of water in this layer (and up to
the 3-rd and 4-th hydration layers), thus slowing down the
bacterial cell movement. In addition, trehalose self-associates in
aqueous solution in a concentration dependent manner to form
clustering networks, affecting the dynamic properties of the
solution. Through extensive solvation, trehalose has a potent
ability to restructure water in the solution and enhance the
hydrogen bonding between water molecules, thus contributing to the
gelation phenomena and the biofilm formation.
During the next stage of the biofilm development (the formation of
bacterial colonies and the maturation of biofilm), the bacteria
will continuously produce trehalose as a general stress response
metabolite and an osmoprotectant in response to constantly varying
environmental conditions, such as increased cell density, nutrients
limitations, and waste products accumulation in the biofilm. Then,
the continuous trehalose--water interactions, with attraction of
new water molecules and further restructuring of water, will result
in formation of new layers of the biofilm and gradually increased
biofilm volume. During this stage, bacteria will release into the
biofilm matrix various extracellular substances, including specific
proteins (adhesins, matrix interacting factors), compatible
solutes, signaling molecules, metabolic end- or by-products, such
as polysaccharides, lipids, phospholipids, and the detritus from
aging and lysed cells, which will contribute to the formation of
the tertiary structure of the biofilm, stabilization of the biofilm
architecture, thickening of the biofilm matrix, and increased
density of the biofilm.
During the late stage (the dispersal stage of microbial biofilm),
as the biofilm ages, the amount of trehalose in the superficial
layers of the biofilm will gradually decrease due to higher
accumulation of trehalose in the deeper layers adjacent to the
microbial cells, so that the trehalose restructuring effect on
water, the strengthening effect on the hydrogen bonds between water
molecules, and the aggregation forces between the microbial cells
will gradually diminish and favor the sloughing off of the
superficial layers of the biofilm, containing the pathogenic
microorganisms that already have been exposed to the host defense
factors and various antimicrobial substances (if used). These
microorganisms, disseminating from the sloughing off superficial
layers of the "parenting biofilm", will be covered by the thin
layer of their "parenting" matrix gel, or just covered by
superficial water layer(s) formed by their own trehalose for
protection from osmotic stress during their move through the body
tissues and fluids. These disseminating microorganisms may express
higher resistance to the host defense factors and antimicrobials
(due to their previous exposure to higher concentrations of
antimicrobials in the superficial layers of"parenting biofilm") and
potentially increased virulence, so that the newly formed
disseminated biofilm sites, containing these pathogenic
microorganisms, will contribute to further increase in
antimicrobial resistance and virulence in the initial
microorganisms forming the "initial biofilms".
At any stage of the biofilm development, the microorganisms
embedded in the biofilm matrix will respond to any environmental
assault on the biofilm, including the use of various harmful
substances (i.e. antimicrobials, disinfectants, and various other
anti-biofilm compounds) by additional production of trehalose as a
general stress response metabolite and an osmoprotectant, that will
result in further increase in the biofilm gel matrix volume and
density, thus preventing the penetration of harmful substances into
the biofilm and protecting the microorganisms from killing.
Trehalose was detected along with other sugars, di-, oligo-, and
polysaccharides in the laboratory-grown microbial biofilms, in the
research studies mostly aimed at either evaluating the effect of
various nutrients on microbial biofilm formation, or analyzing the
content of the biofilm exopolymer matrix.
For example, trehalose was detected in a small amount (3%), along
with glycerol (5%), mannitol (18%), and glucose (74%), in the
monosaccharide-polyol fraction of the aerial-grown hyphae of the
Aspergillus fumigatus biofilm; all hexoses and polyols were found
intracellularly in the same proportion as extracellularly (Beauvais
A, Schmidt C, Guadagnini S, Roux P, Perret E, Henry C, Paris S,
Mallet A, Prevost M, and Latge J P, "An extracellular matrix glues
together the aerial-grown hyphae of Aspergillus fumigates,"
Cellular Microbiology, 2007; 9(6): 1588-1600.). In another example,
biofilm development on stainless steel by Listeria monocytogenes
(the most common biofilm-producing pathogen in the food industry),
was enhanced by the presence of mannose or trehalose as nutrients
in the growth media, with trehalose being superior to mannose in
constant biofilm production during 12 days of incubation at 21
degrees C. (Kim K Y and Frank J F, "Effect of nutrients on biofilm
formation by Listeria monocytogenes on stainless steel," Journal of
food protection, 1995; 58(1): 24-28.). In another study, the
formation of a structurally and metabolically distinctive biofilm
by Streptococcus mutans (the most common pathogen in dental
biofilms), was enhanced by the combination of sucrose and starch,
compared with sucrose alone, in the presence of surface-adsorbed
salivary a-amylase and bacterial glucosyltransferases, with
upregulation of genes associated with maltose uptake/transport and
fermentation/glycolysis (Klein M I, DeBaz L, Agidi S, Lee H, Xie G,
Lin A H, Hamaker B R, Lemos J A, and Koo H, "Dynamics of
Streptococcus mutans Transcriptome in Response to Starch and
Sucrose during Biofilm Development," PLoS ONE, 2010; 5(10): 1-13.).
In the next study, the yeasts from hydrocarbon-polluted alpine
habitats (Cryptococcus terreus--strain PB4, and Rhodotorula
creatinivora--strains PB7 and PB12) synthesized and accumulated
glycogen (both acid- and alkali-soluble) and trehalose during
growth in culture media, containing either glucose or phenol as a
sole carbon and energy source, with higher biofilm formation by
both strains of Rhodotorula creatinivora (Krallish I, Gonta S,
Savenkova L, Bergauer P, and Margesin R, "Phenol degradation by
immobilized cold-adapted yeast strains of Cryptococcus terreus and
Rhodotorula creatinivora," Extremophiles, 2006; 10(5):
441-449.).
In contrast to the previous results, the laboratory-grown wild type
Enterococcus faecalis formed strong biofilm in the presence of
maltose or glucose in the growth media, and formed very little
amount of biofilm in medium containing trehalose (Creti R, Koch S,
Fabretti F, Baldassarri L, and Johannes H, "Enterococcal
colonization of the gastro-intestinal tract: role of biofilm and
environmental oligosaccharides," BMC Microbiology, 2006; 6:
660-668.).
Since trehalose is the most abundant disaccharide in yeasts and
fungi, the biofilm matrix of any biofilm-based yeast or fungal
infections, and/or multispecies biofilms which include yeasts
and/or fungi, can be more resistant to penetration by
antimicrobials.
Enzyme trehalase (.alpha.,.alpha.-trehalase;
.alpha.,.alpha.-trehalose-1-C-glucohydrolase, EC 3.2.1.28)
Enzyme trehalase (a, a-trehalase), highly specific for the
non-reducing disaccharide trehalose, directly degrades trehalose
into two molecules of glucose on hydrolysis, being this the only
trehalose degradation pathway reported up to today. Trehalase
phylogeny unveiled three major branches comprising those from
bacteria, plant and animals, and those from fungal origin.
Crystallographic study of bacterial trehalase indicated that this
enzyme's structures are highly conserved in spite of the marked
differences found at the sequence level, suggesting a bacterial
origin for the trehalases in contrast to an eukaryotic origin, as
previously proposed (Barraza A, Sanchez F. "Trehalases: A neglected
carbon metabolism regulator?", Plant Signal Behva 2013; 8:
e24778.). Trehalose degradation by trehalase appears to be
important, perhaps essential, in the life functions of various
lower organisms, including yeasts, fungi, bacteria, insects, and
invertebrates (Nwaka S and Holzer H, "Molecular biology of
trehalose and trehalases in the yeast, Saccharomyces cerevisiae,"
Prog. Nucleic Acid Res. Mol. Biol., 1998; 58: 197-237.). Enzyme
trehalase also has been reported to be present in many
macroorganisms (including mammals--animals and humans) and vascular
plants, but the functions and properties of this enzyme were not
fully elucidated (Elbein A D, "The metabolism of
.alpha.,.alpha.-trehalose," Adv. Carbohyd. Chem. Biochem, 1974; 30:
227-256.); (Elbein A D, Pan Y T, Pastuszak I, and Carroll D, "New
insights on trehalose: a multifunctional molecule," Glycobiology,
2003; Vol. 13, No 4: 17R-27R.).
As many as 541 model variants of trehalase can be found in the
Protein Model Portal (http://www.proteinmodelportal.org/). A few of
these models corresponding to different enzyme variants
(isoenzymes) are shown in FIGS. 2a through 2d.
In lower forms of life (yeasts, fungi, bacteria), there are two
main types of trehalase enzyme: neutral trehalase (NTH) and acid
trehalase (ATH), which are encoded by two different genes. Most of
the trehalase activity in these microorganisms, comes from the
neutral trehalase, located in the cytosol, with the pH optimum of
about 7, highly specific for trehalose as the substrate, and
inactive on cellobiose, maltose, lactose, sucrose, raffinose, and
mellibiose; this enzyme has also a specific regulatory function
(App H and Holzer H, "Purification and characterization of neutral
trehalase from the yeast ABYS1 mutant," J. Biol. Chem., 1989; 264:
17583-17588.). The acid or vacuolar trehalase has a pH optimum of
4.5 and is also very specific for trehalose as the substrate,
showing no activity with cellobiose, maltose, lactose, sucrose, and
mellibiose; this enzyme acts in the periplasmic space where it
binds exogenous trehalose to internalize it for further cleavage it
in the vacuoles to produce free glucose (Mittenbuhler K and Holzer
H, "Purification and characterization of acid trehalases from the
yeast SUC2 mutant," J. Biol. Chem., 1988; 263: 8537-8543.);
(Stambuk B U, de Arujo P S, Panek A D, and Serrano R, "Kinetics and
energetics of trehalose transport in Saccharomyces cerevisiae,"
Eur. J. Biochem., 1996; 237: 876-881.).
The activities of both trehalases (NTH and ATH) are low in yeast
cells growing exponentially, but high during stationary phase
growth after glucose has been depleted (Winkler K, Kienle I,
Burgert M, Wagner J C, and Holzer H, "Metabolic regulation of the
trehalose content of vegetative yeast," FEBS Lett., 1991; 291:
262-272.). ATHI deletion mutant of the yeast S. cerevisiae cannot
grow in the medium with trehalose as the carbon source, but a
Candida utils mutant strain is able to utilize extracellular
trehalose as carbon source despite of the lack of ATH activity.
Various bacteria, such as E. coli, have trehalases that also may
supply exogenous trehalose and glucose via the phospho-transferrase
system (PTS) (Horlacher R, Uhland K, Klein W, Erhmann M, and Boos
W, "Characterization of a cytoplasmic trehalase of Escherichia
coli," J. Bacteriol., 1996; 178: 625-627.).
In the plant kingdom, enzyme trehalase is ubiquitous, being
involved in carbon metabolism in both lower and higher plants.
Although sugar trehalose is rare in higher (vascular) plants, it
has been demonstrated that trehalase in these plants could take a
part in the degradation of trehalose derived from the
plant-associated bacteria in symbiotic interactions; it has been
also suggested that trehalase in higher plants could play
additional role in the defense against parasites and other
pathogenic organisms (in plant--pathogen interactions) (Muller J,
Wiemken A, and Aeschbacher R, "Trehalose metabolism in sugar
sensing and plant development," Plant Sci., 1999; 147: 37-47.);
(Muller J, Aeschbacher R A, Wingler A, Boller T, and Wiemken A,
"Trehalose and trehalase in Arabidopsis," Plant Physiol., 2001;
125: 1086-1093.).
Though disaccharide trehalose is not produced in mammals, the
enzyme trehalase exists in mammals (including humans) in the kidney
brush border membranes and in the intestinal villi membranes; the
role of trehalase in kidney is still not clear, but in the
intestine its function is to hydrolyze ingested trehalose
(Dahlqvist A, "Assay of intestinal disaccharidases," Anal.
Biochem., 1968; 22: 99-107.); (Ruf J, Wacker H, James P, Maffia M,
Seiler P, Galand G, Kiekebusch A, Semenza G, and Mantei N, "Rabbit
small intestine trehalase. Purification, cDNA cloning, expression
and verification of GPI-anchoring," J. Biol. Chem, 1990; 265:
15034-15040.); (Yonemaya Y and Lever J E, "Apical trehalase
expression associated with cell patterning after inducer treatment
of LLC-PK monolayers," J. Cell. Physiol., 1987; 131: 330-341.).
Only one type of trehalse (.alpha.,.alpha.-trehalase, highly
specific enzyme for direct degradation of trehalose into two
molecules of glucose) is present in humans. Trehalase produced by
the glands of Lieberkuhni in the small intestine is a constituent
of the intestinal juice along with other specific saccharidases,
such as maltase, sucrase-isomaltase complex, and
Beta-glycosidase-lactase (Mayes P A, "Carbohydrates of physiologic
significance," In: Harper's Biochemistry, 25th ed, 2000, pp.
149-159, Appleton & Lange, Stamford, Conn.); (Rodwell V W and
Kennelly P J, "Enzymes: General Properties; Enzymes: Kinetics," In:
Harper's Biochemistry, 25th ed, 2000, pp. 74-102, Appleton &
Lange, Stamford, Conn.). As with all other disaccharidases,
trehalase remains attached to the brush border of the enterocyte in
the intestinal lumen while the catalytic domain is free to react
with the substrate. There is little free trehalase activity in the
intestinal lumen; most activity is associated with small "knobs" on
the brush border of the intestinal epithelial cells. A small
fraction (approximately 0.5%) may be absorbed by passive diffusion,
as shown for other disaccharides, in patients with trehalase
deficiency (van Elburg R M, Uil J J, Kokke F T M, Mulder A M, van
dr Broek W G M, Mulder C J J, and Heymans H S A, "Repeatability of
the sugar-absorption test, using lactulose and mannitol, for
measuring intestinal permeability for sugars," J. Pediatr.
Gastroenterol. Nutr., 1995; 20: 184-188.). Trehalase activity have
been found also in the renal cortex, plasma, urine, liver and bile,
although function of the enzyme in these locations is not clear
yet; it is likely that trehalase in the urine and bile can be
incidental to its presence in the kidney and liver (Eze L C,
"Plasma trehalase activity and diabetes mellitus," Biochem Gen.,
1989; 27: 487-495.). A complete cDNA clone encoding human trehalase
(a protein of 583 amino acids with calculated molecular weight of
66 595 kDa), a glycoprotein of brush-border membranes, has been
isolated from a human kidney library; the deduced amino acid
sequence of the human trehalase enzyme showed similarity to
sequences of the enzyme trehalase from rabbit (81%), silk worm,
Tenebrio molitor (43%), Escherichia coli (33%); human trehalase
also resembled yeast acidic trehalase--ATH (25% identity) and
neutral trehalase--NTH (26.5% identity); by homology with mammalian
trehalase from other species, human enzyme is an ectoenzyme whose
hydrophobic region at the carboxyl terminus is linked to the plasma
membrane by GPI anchor (Reiko Ishihara, Shigeru Taketani, Misa
Sasai-Takedatsu, Minoru Kino, Rikio Tokunaga, Yohnosuke Lobayashi,
"Molecular cloning, sequencing and expression of cDNA encoding
human trehalase", Cene 202 (1997) 69-74.). The research data showed
that mammalian trehalase is encoded by a single gene and is
probably expressed as one form in various tissues (Ruf, J., et.
al., 1990, "Rabbit small intestinal trehalase: purification, eDNA
cloning, expression, and verification of
glucosylphosphatidylinositol anchoring", J. Biol. Chem. 265,
15034-15039; Reiko Ishihara, et. al., "Molecular cloning,
sequencing and expression of cDNA encoding human trehalase", Gene
202 (1997) 69-74.
Biochemical properties of the human enzyme
.alpha.,.alpha.-trehalase include: high specificity for the
substrate (disaccharide trehalose--a, a-trehalose) method of
activation--direct contact with the substrate (trehalose) optimal
conditions for activity--in the range of pH between 5.0 and 7.0
(similar to the other disaccharidases) end product of action--2
molecules of D-glucose heat sensitivity--as a glycosylated protein
is probably similar to the other disaccharidases catalytic
efficiency--high due to the high specificity for the substrate
trehalose coenzymes or metal ions for activity--not needed
co-variants of enzyme--unknown
The main function of human .alpha.,.alpha.-trehalase is to
hydrolyze ingested disaccharide trehalose into glucose. Trehalase
deficiency is a known metabolic condition in humans, when the body
is not able to degrade disaccharide trehalose into two molecules of
glucose and digest it. People with enzyme trehalase deficiency
experience vomiting, abdominal discomfort and diarrhea after eating
mushrooms (rich in trehalose) of any other food containing
trehalose. Trehalase enzyme deficiency in most cases appear to be
inherited in an autosomal recessive manner (Kleinman R E, Goulet O,
Mieli-Vergani G, Sherman P M, In: Walker's Pediatric
Gastrointestinal Disease: Physiology, Diagnosis, Management, 5-th
edition, 2008); (Semenza, G., Auricchio, S., and Mantei, N. In: The
Metabolic & Molecular Bases of Inherited Disease; 8-th ed.,
2001; Chapter 75: Small Intestinal Disaccharidoses. McGraw-Hill,
New York.). Isolated intestinal trehalase deficiency is found in
approximately 8% of Greenlanders; it is not infrequent among Finns,
but is believed to be rare elsewhere. The low (2%) incidence of
isolated trehalase enzyme deficiency was described in the
populations from the USA, UK, and mainland Europe (Bergoz R,
Valloton M C, and Loizeau E, "Trehalase deficiency," Ann. Nutr.
Metab., 1982; 26: 191-195.). In the UK, from 400 patients
investigated for suspected malabsorption, 369 (92%) had normal
intestinal histology on biopsy, with the normal range of trehalase
at 4, 79-37, 12 U/g protein; 31 (8%) patients with villous atrophy
had a diagnosis of coeliac disease and significantly reduced
activity of disaccharidases, including trehalase, with recovered
function of all enzymes (except lactase) after treatment with a
gluten-free diet; the authors concluded that there is no basis for
routine determination of trehalase activity in the population of
the UK (Murray I A, Coupland K, Smith J A, Ansell I D, Long R G,
"Intestinal trehalase in a UK population: Establishing a normal
range and the effect of disease," Br. J. Nutr., 2000; 83(3):
241-245.). In Belgium, in intestinal biopsy samples from 200
patients with abdominal symptoms and diarrhea, total
.alpha..alpha..alpha.-trehalase deficiency (0-12 U/g mucosa) was
detected in 18 (9%) cases, partial deficiency (3-12 U/g mucosa)--in
39 (19.5%) cases, and only 4 patients (2%) presented selective
.alpha..alpha..alpha.-trehalase deficiency with otherwise normal
other disaccharidases; these data suggested that
.alpha..alpha..alpha.-trehalase deficiency can be more common than
it is believed (Buts J P, Stilmant C, Bernasconi P, Neirinck C, De
Keyser N, "Characterization of alpha, alpha-trehalase released in
the intestinal lumen by the probiotic Saccharomyces boulardii,"
Scandinavian Journal of Gastroenterology, 2008; 43 (12):
1489-1496.).
The importance of enzyme trehalase was demonstrated in certain
pathologic conditions, including birth defects and genetic
abnormalities: low or absent intestinal trehalase was detected in
the sample of amniotic fluid from a fetus with anal imperforation,
whereas a higher than normal level of renal trehalase activity was
found in amniotic fluid from a fetus with polycystic kidney disease
(Elsliger M A, Dallaire L, Potier M, "Fetal intestinal and renal
origins of trehalase activity in human amniotic fluid," Clin Chim
Acta, Jul. 16, 1993; 216(1-2): 91-102.). Also, low intestinal
trehalase enzyme level was detected in amniotic fluid on
amniocentesis in 14 pregnant women at 1 in 4 risk for a child with
cystic fibrosis, screened at the 18-th week of gestation; and in
two terminated at the 19-th week cases, histochemical lesions
characteristic of cystic fibrosis were seen in exocrine glands,
including the pancreas and intestinal mucosa of both fetuses, and
the total protein content in the meconium of these fetuses was also
significantly higher than in the controls (Szabo M, Teichmann F,
Szeifert G T, Toth M, Toth Z, Torok O, Papp Z, "Prenatal diagnosis
of cystic fibrosis by trehalase enzyme assay in amniotic fluid,"
Article first published online: 23 Apr. 2008; DOI: 10.1111/j.
1300-0004. 1985.tb01211.x.). The trehalase enzyme assay in amniotic
fluid was recommended as a genetic test for prenatal diagnosis of
cystic fibrosis. The latest genetic studies in 2942 full-heritage
Pima Indians and 3897 "mixed" heritage Native Americans with
Diabetes type 2 (T2D), found strong correlation with trehalase
enzyme activity in plasma of people with T2D; four single
nucleotide polymorphisms (SNPs) were detected in their trehalase
gene (TREH) that were associated with T2D (Yunhua L Muller, Robert
L Hanson, William C Knowler, Jamie Fleming, Jayita Goswani, Ke
Huang, Michael Traurig, Jeff Sutherland, Chris Wiedrich, Kim
Wiedrich, Darin Mahkee, Vicky Ossowski, Sayuko Kobes, Clifton
Bogardus, Leslie J Baier, "Identification of genetic variation that
determines human trehalase activity and its association with type 2
diabetes", Humangenetik 2013-06-01.). The Chinese research studies
on genetic risk factors for glioma (one of the most aggressive
human tumors), indicated that more than 100 single nucleotide
polymorphism (SNPs) are associated with the risk of glioma,
including the SNPs in trehalase (TREH) gene, and provided evidence
for three glioma susceptibility genes--TREH, IL4, and CCDC26
(Shangu Li, Tianbo Jin, Jiayi Zhang, Huiling Lou, Bo Yang, Yang Li,
Chao Chen, Yongsheng Zhang, Shanqu Li, Tianbo Jin, Jiayi Zhang,
Huiling Lou, Bo Yang, Yang Li, Chao Chen, Yongsheng Zhang,
"Polymorphisms of TREH, IL4 and CCDC26 genes associated with risk
of glioma", Cancer Epidemiology 2012-06-01.). The correlation
between intestinal histology and trehalase activities during
intestinal injury has been shown in clinical studies in patients
with intestinal ischemia-reperfusion injury (Stefan Toth, Timea
Pekarova, Jan Varga, Vladimira Tomeckova, Stephan Toth, Lucia
Lakyova, Jarmila Vesela, "Trehalase as a possible marker of
intestinal ischemia-reperfusion injury", Acta Biochimica Polonica
2013-01-01.).
Since ingestion of large quantities of foods containing trehalose
is not common worldwide, the real frequency of trehalase deficiency
in various populations around the world is mostly unknown. However,
it should be noted, that over the last two decades, in addition to
natural sources of trehalose in the food (mostly, mushrooms, algae,
baker's yeasts), it has been approved in some countries, including
the USA, as an additive in the preparation of dried, frozen, and
processed food, and as a moisture retainer in various products
(including ice cream, and baked goods), with no requirements for
labeling of this constituent in prepared food or other products
(Abbott P J and Chen J, WHO Food Additives Series 46: Trehalose.
International Programme on Chemical Safety. Accessed Feb. 4, 2010,
available at:
http://www.inchem.org/documents/jecfa/jecmono/v46je05.htm.).
The amount of enzyme trehalase normally produced for digestion and
utilization of exogenous trehalose is appropriate for healthy
people, but is far less than what is needed for people with
biofilm-based chronic infections, especially for individuals with
trehalase enzyme deficiency. Therefore, the use of enzyme
trehalase, along with other enzyme formulations and antimicrobials
(including antibiotics), can greatly enhance the effectiveness of
various treatment protocols for biofilm-based chronic
infections.
Therefore, at least one basis for the presently disclosed
compositions and methods is the addition of enzyme trehalase,
highly specific to the hydrolysis of the trehalose constituent of
microbial biofilms, to treatment protocols for biofilm-based
chronic infections in order to increase the effectiveness of
existing treatment modalities.
To avoid possible immunogenicity and/or toxicity of any other
sources of trehalase (animal-derived or microbial-derived enzymes)
while used in humans, the best source of trehalase for use in
humans can be a human genetic recombinant enzyme (human trehalase
gene script expressed in rice genetically modified for human enzyme
production); this manufacturing method will be analogous to
production of synthetic human enzyme Lysosyme ("Lysobac") produced
by Sigma-Aldrich, USA. Also, plant-derived trehalase (sugar cane
trehalase--.alpha.,.alpha.-trehalase, that is located in the
cytosol of the plant cells), can be used in humans; in anecdotal
cases of using natural row sugar cane sticks for chewing as a sweet
treat (substitute for chewing gum and candy) and using a raw
(non-processed) fresh pressed sugar cane juice for food (as sugar
substitute), it was demonstrated that the adult and children
population in remote regions of Central America had healthy teeth,
absence of dental caries and periodontal diseases, and healthy
condition of the epithelial lining of the oral cavity (personal
observations, not published research data). So that, the enzyme
trehalase in the raw sugar cane juice (sugar cane
trehalase--.alpha.,.alpha.-trehalase,) seems to be safe for use in
humans, and can be used in combination with antimicrobials for
prevention and treatment of microbial biofilm in the oral cavity.
Also, the process of manufacturing of sugar cane trehalase can be
cost-efficient regarding its possible wide use in medical and
health care fields. Trehalse can be manufactured in various forms
(powder, liquid, gel, tablets, and capsules) that can be tailored
to specific applications in humans and delivered to any specific
location in the body where biofilm is the issue (mostly mucosal
linings of the oral cavity, GI tract, respiratory tract, and
urinary tract, and skin lesions and wounds); trehalase can be used
in combination with other saccharidase enzymes (dextranase and
lyase), included in existing formulations of digestive enzymes,
included in various existing formulations of vitamins, used in
combination with various natural substances (anti-inflammatory and
immune-modulating compounds and anti-oxidants), and can be used in
combination with various antibiotics and natural antimicrobials
(antimicrobial peptides of human- or plant-origin) to treat
biofilm-based infections in a human body and to control biofilms on
medical devices, medical equipment, and various medical implants.
For industrial applications (water, gas, and oil pipelines, HV/AC
systems), trehalase can be obtained from natural sources (plants,
algae, fungi) using known manufacturing methods (for example, the
U.S. Pat. No. 5,593,869 Jan. 14, 1997 "Method of manufacturing
sugars by trehalase" includes the description of manufacturing
novel enzyme trehalase from green algae of Lobosphaera,
Chlorellaceae, Chlorofyceae, Chlorophyta families); in another
example--method of extraction of trehalase from Sugar Cane is
described in PH Dissertation of Susan Bosch, 2005, "Trehalose and
Carbon Partitioning in Sugar Cane, Chapter 5, pp. 123-146,
University of Stellenbosch, South Africa). However, to date, no
available medical/health scientific information shows evidence of
enzyme Trehalase as a component of any prescription drugs, natural
antimicrobials preparation and their combinations, OTC products,
nutritional supplements, vitamins combinations, digestive enzymes
formulations and/or enzymatic formulations used for local treatment
of microbial biofilms on skin and mucosal lining surfaces of
humans, or as a component of systemic enzymes formulations, or as a
component of compositions and methods for biofilm treatment on
medical devices, implants, and equipment, or being a component of
various disinfecting combinations for prevention and treatment of
microbial biofilm on various surfaces in hospitals (including
surgical units), medical offices and other medical/health settings,
and public places. Also, there is no available information about
the enzyme trehalase being a component of any existing anti-biofilm
formulations and/or any methods for microbial biofilm prevention
and treatment in industrial fluid conduits (water, gas, oil
pipelines), and various industrial and household equipment.
Embodiments for the Treatment of Biofilm-Based Infections
To increase effectiveness of existing treatment modalities and
protocols for biofilm-based chronic infections in the human body,
combination of trehalase with antimicrobials can be used alone or
in combination with other enzymes and substances (antioxidants,
anti-inflammatory and immune-modulating), being included in various
formulations and methods of their use, for direct application to
the sites of infectious biofilm (directly accessible mucosal
linings of the respiratory tract, GI-tract, genito-urinary tract,
eyes, skin, open wounds, etc.) and/or for systemic use to treat
biofilm-based infections in directly inaccessible (or hardly
accessible) sites of infection and in the bloodstream.
Simultaneously, trehalase can be used in combination with natural
substances (natural antimicrobials, human antimicrobial peptides,
anti-oxidants, anti-inflammatory and immune-modulating substances)
to support and/or enhance body's own abilities to prevent microbial
biofilm formation or degrade formed biofilm, to kill invading
pathogens and eliminate them from the body, to restore normal
homeostasis and immune system function, and to repair any tissue
damages resulted from biofilm-based infections.
In direct application to the sites of microbial biofilm in a human
body, trehalase enzyme could be used as a component of formulations
of hydrolytic enzymes (including dextranase and alginate-lyase) and
antimicrobilals, applied in a single step to the site of infectious
biofilm; or trehalase can be included in anti-biofilm combinations
of enzymes and antimicrobials used in a multi-step procedure,
starting as the first (pretreatment) step with application of
combination of hydrolytic enzymes (trehalase, dextranase,
alginate-lyase) with an exposition time sufficient to initiate
degradation of the biofilm matrix, followed by the second step with
application of combination of proteolytic, fibrinolytic, and
lipolytic enzymes over a corresponding appropriate exposition time
to further degrade the biofilm matrix, followed in a third step by
application of antimicrobials (or their combinations) specific for
the infection(s) involved, or polymicrobial antibiotics, in
combination with trehalase for prolonged use. In this 3-step
method, the initial degrading effect on biofilm matrix (provided by
trehalase in combination with other hydrolytic enzymes), will ease
and potentiate the action of proteolytic, fibrinolytic, and
lipolytic enzymes to further degrade the biofilm matrix and provide
access to microbial cells for antimicrobal substances combined with
trehalase to gradually kill infectious pathogens and prevent the
Biofilm Induction Response (BIR) to sub-MIC doses of antimicrobials
at the initiation of treatment course. The BIR is a well-known
phenomenon that is of concern in clinical practice, when killing
concentration of antibiotics gradually increases at the site of
microbial biofilm on initiation of treatment, allowing bacteria to
initiate BIR at the very initial time of treatment (within minutes
to a few hours). In addition, at the end of treatment course, some
dormant microorganisms (so called persister cells) who survived
antimicrobial treatment, will come back to active growing state
from dormancy and will form biofilm as BIR, being exposed to the
decreasing dose of antimicrobials (down to sub-MIC dose) (Jeffrey
B. Kaplan, "Antibiotic-Induced Biofilm Formation. Review; Int J
Artif Organs 2011; 34(9): 737-751.). Therefore, the presence of
trehalase in combination with antimicrobials during full treatment
course, will increase the effectiveness of used antimicrobials and
possibly reduce antimicrobial resistance in biofilm-forming
infectious pathogens.
As a systemic enzyme, trehalase in combination with other
saccharidases should be used as a time-delayed release
substance(s), or being included in multi-enzyme formulations with
time-delayed release of constituent(s) to avoid early degradation
of trehalase and other hydrolytic enzymes (dextranase and
alginate-lyase) by proteolytic enzymes in the upper GI tract
(stomach and duodenum) and/or by proteolytic enzymes in
administered formulations, and finally being released in the small
intestine for further absorption. In this way, trehalase can be
supplied for direct absorption and distribution via the bloodstream
to hardly accessible "niches" of biofilm-based infections, for
example, on the inner lining of the blood vessels, in bones,
joints, on medical devices and implants).
The length of the time for delayed release can be established for
trehalase in combination with other hydrolytic enzymes (dextranase
and alginate-lyase), so that the release of these enzymes
combination will occur in the small intestine, being protected from
proteolytic enzymes in the upper GI tract (stomach and duodenum).
Also, differential time delays can be established for combination
of trehalase with other hydrolytic enzymes (dextranase and
alginate-lyase) and any co-administered proteolytic enzymes (having
different pH for their proteolytic activities) or a combination of
these proteolytic enzymes, to avoid deleterious action of
proteolytic compounds on trehalase and other hydrolytic enzymes
(dextranase and alginate-lyase). In conventional digestive or
systemic enzyme formulations currently on the market, the contained
hydrolytic enzymes (di-, oligo-, and polysaccharidases) typically
are not protected from deleterious action of various proteolytic
enzymes included in existing formulations.
Upper Respiratory Tract
The major biofilm-forming species of pathogens affecting the upper
respiratory tract include Haemophilus influenzae, Klebsiella
pneumoniae, Pneumococcus, Streptococcus spp., Staphylococcus spp.,
Pseudomonas aeruginosa, Candida spp., and Aspergillus spp. For
local treatment of biofilm-based infections in the upper
respiratory tract (chronic sinusitis, rhinosinusitis, tonsillitis,
pharyngitis, and otitis media), trehalase enzyme can be used in
combination with other enzymes, antimicrobials specific to the
present pathogens or polymicrobial antibiotics, or natural
antimicrobials, and anti-inflammatory and immune-modulating
substances in direct application to the sites of infectious biofilm
on mucosal lining, in liquid form as a saline-based solution for
instillations, irrigations, and sprays, as well as in gel,
ointment, and powder forms.
In direct application to the sites of infectious biofilm on mucosal
lining of the upper respiratory tract, trehalase enzyme could be
used as a component of formulations of hydrolytic enzymes (di-,
oligo-, and polysaccharidases, including dextranase and
alginate-lyase as most frequently used hydrolytic enzymes) and
antimicrobilals, applied in a single step to the site of infectious
biofilm; or trehalase can be included in anti-biofilm combinations
of various enzymes, antimicrobials, anti-inflammatory and
immune-modulating substances, and used in a multi-step procedure,
starting as the first (pretreatment) step with application of
combination of hydrolytic enzymes (trehalase, dextranase,
alginate-lyase) with an exposition time sufficient to initiate
degradation of the biofilm matrix, followed by the second step with
application of combination of proteolytic, fibrinolytic, and
lipolytic enzymes over a corresponding appropriate exposition time
to further degrade the biofilm matrix, followed in a third step by
application of antimicrobials (or their combinations) specific for
the infection(s) involved, or polymicrobial antibiotics, in
combination with trehalase, anti-inflammatory and immune-modulating
substances for prolonged use. Local treatment can be reinforced by
systemic use (introduced via GI tract for absorption and
distribution via blood stream to the sites of infectious biofilm)
of multi-enzyme formulations (including trehalase in a time-delayed
release form), anti-inflammatory and immune-modulating substances,
along with systemic use of antibiotics preferably with
polymicrobial activity.
For local treatment of Pseudomonas aeruginosa infectious biofilm on
mucosal lining of the upper respiratory tract, the enzyme
alginate-lyase (highly specific for degradation of the
polysaccharide alginate--an important constituent of Pseudomonas
aeruginosa biofilm), should be also added to combination of
trehalase with antibiotics, anti-inflammatory and immune-modulating
substances for prolonged use as the third step of local
application. For Streptococcus spp. infections, the enzyme
dextranase (highly specific for degradation of the
dextrans--oligosaccharides produced by Streptococcus spp., which
facilitate microbial adhesion to the mucosal surfaces and biofilm
formation, should also be added to combination of trehalase with
antibiotics, anti-inflammatory and immune-modulating substances for
prolonged use as the third step of treatment in local application
to treat microbial biofilm in the upper respiratory tract.
Otitis Media
For otitis media with or without effusion, treatment should include
a systemic enzymes formulation (including trehalase in combination
with other saccharidases in time-delayed release form) along with
systemic antibiotics. The systemic treatment can be reinforced with
local treatment applied to the lining of the nasal cavity to
address possible spread of infection to the middle ear from the
nasal and sinus cavities; this local treatment should include
combination of trehalase with anti-inflammatory and
immune-modulating substances, and polymicrobial antibiotics or
natural antimicrobial substances with wide range of anti-infectious
activity (for example, colloidal silver spray). For specific
treatment of Pseudomonas aeruginosa and/or Streptococcal biofilms,
the enzymes alginate-lyase and dextranase should be added to
combination of trehalase with antimicrobials. Delivery of the local
treatment to the inner ear can be done by a nasal instillation with
a pathway through the Eustachian tube into the middle ear.
For otitis media with effusion and installed tympanic tubes, the
abovementioned systemic and local treatments should be reinforced
by an additional step: the installed tympanic tubes can be covered
inside with trehalase in combination with polymicrobial antibiotics
or natural antimicrobials with wide range of antimicrobial
activities.
Lower Respiratory Tract
Treatment of biofilm-based infections in the lower respiratory
tract, should include: a) the use of systemic enzymes (with
trehalase and other saccharidases in time-delayed release form)
along with systemic antibiotics; b) local treatment using
bronchi-alveolar or whole lung lavage in a multi-step procedure,
including as the first step--the use of trehalase in combination
with other saccharidases (alginate-lyase, dextranase) in a
saline-based solution, followed by proteolytic, fibrinolytic, and
lipolytic enzymes in a saline-based solution as the second step,
and antibiotics in combination with trehalase in a saline-based
solution as the third step; c) nasal instillation of trehalase in
combination with anti-inflammatory and immune-modulating
substances, and polymicrobial antibiotics or natural antimicrobial
substances with wide range of antimicrobial activities.
Additional contributing factors to chronic biofilm-based infectious
conditions are: genetic trehalase enzyme deficiency (a rare genetic
disease listed by NIH Genetic and Rare Diseases Information
Center), genetic trehalase enzyme deficiency in individuals with
cystic fibrosis, and artificial trehalase deficiency due to
widespread use of trehalose in the food industry as an approved
additive in the preparation of dried and frozen foods, and as a
moisture conservant, in various foods, such as an ice cream and
baked goods.
Taking into account genetic trehalase deficiency in cystic fibrosis
patients, uncontrolled consumption of trehalose in food is a
favorable factor for thick biofilm formation on the mucosal lining
of the upper and lower respiratory tracts in such individuals.
Pseudomonas aeruginosa, in symbiosis with other bacteria and fungi,
exploits this environment, creating thick polymicrobial biofilm
which is almost impossible to eradicate with long-term antibiotic
therapy alone (although such therapy can support the patient).
Including the enzyme trehalase in combinations with antibiotics,
anti-oxidants, anti-inflammatory and immune-modulating substances
in various protocols for local and systemic treatments of
biofilm-based chronic infections in patients with cystic fibrosis,
will significantly increase positive outcomes of such treatment
protocols and improve patients' quality of life. To address this
thick polymicrobial biofilm at any stages of its development,
treatment should include: a) the use of systemic enzymes (with
trehalase and other saccharidases in time-delayed release form),
along with systemic antibiotics; b) local treatment using
bronchi-alveolar or whole lung lavage in a multi-step procedure,
including as the first step--the use of trehalase in combination
with other saccharidases (alginate-lyase, dextranase) in a
saline-based solution, followed by proteolytic, fibrinolytic, and
lipolytic enzymes in a saline-based solution as the second step,
and antibiotics in combination with trehalase in a saline-based
solution as the third step; c) using special enzyme "Pulmozyme" to
thin the mucus in the airways of CF patients. This treatment can be
reinforced by using nasal instillation of trehalase in combination
with anti-inflammatory and immune-modulating substances, and
polymicrobial antibiotics or natural antimicrobials with wide range
of antimicrobial activities.
Native Valve Endocarditis (NVE), Infectious Endocarditis, and Line
Sepsis
A preferred treatment protocol for NVE, Infectious Endocarditis,
and Line Sepsis as blood stream infections, should include use of
systemic enzyme formulations, with included trehalase and other
saccharidases (preferably, specific to present pathogens) in
time-delayed release form; and trehalase in combination with
antibiotics directed to specific infectious agents, or
polymicrobial antibiotics. The typical organisms involved in these
biofilm-mediated infections include Streptococci spp, Enterococci
spp., Pneumococcus, Staphylococci spp. (both coagulase positive and
negative), gut bacteria, and fungi (most often, Candida albicans
and Aspergillus spp.). Because all these pathogens gain access to
the bloodstream primarily via the oropharynx, GI-tract, and
genito-urinary tract, systemic treatment of NVE, Infectious
Endocarditis, and Line Sepsis should be reinforced by local
treatment of those infections at the sites of origin, including the
previously described multi-step procedure (with application of
trehalase, other enzymes, and antimicrobials) if the sites of
origin represent biofilm-based infections.
Chronic Bacterial Prostatitis (CBP) and Urinary Tract Infections
(UTI)
Use of systemic enzymes with included trehalase and other
saccharidases in time-delayed release form, and antimicrobials in
combination with trehalase will address the presence of
biofilm-based chronic infections in both CBP and UTI. For local
treatment of UTI via bladder instillation, a method after the
fashion of a single-step procedure or the multi-step procedure
disclosed above for treating biofilm on mucosal linings, should be
employed: trehalase in combination with other hydrolytic enzymes
(dextranase, alginate-lyase), and trehalase in combination with
polymicrobial antibiotics and/or natural antimicrobials, can be
applied in a single step to the site of infectious biofilm; or
trehalase can be included in anti-biofilm combinations of enzymes
and antimicrobials used in a multi-step procedure, starting as the
first (pretreatment) step with application of combination of
trehalase with other hydrolytic enzymes (dextranase,
alginate-lyase) with an exposition time sufficient to initiate
degradation of the biofilm matrix, followed by the next
step--application of combination of proteolytic, fibrinolytic, and
lipolytic enzymes over a corresponding appropriate exposition time
to further degrade the biofilm matrix, and in the next
step--application of trehalase in combination with antibiotics (or
their combinations) specific for the present infection(s) involved,
or polymicrobial antibiotics, and/or natural antimicrobials for
prolonged use. For local treatment of CBP, again, trehalase can be
used in combination with other enzymes and antimicrobials in a
single-step application or used in multi-step procedure, but with
higher concentrations of polymicrobial antibiotics and/or natural
antimicrobials delivered directly to the biofilm location within
the prostatic ducts by instillation means (via a medical device
such as a catheter).
GI Tract Infections
GI tract infections are characterized by polymicrobial biofilm
communities along with chronic parasitic and helmintic infections
(nematodes are known to produce and accumulate trehalose). For
treating microbial biofilms in the upper GI tract, trehalase can be
included in combinations of digestive enzymes for fast release,
along with use of trehalase in combination with antibiotics
specific for present infection(s) and/or natural antimicrobials.
For treatment of biofilm-based infections in the lower GI tract,
the formulations of digestive enzymes should include trehalase with
other saccharidases in time-delayed release form to avoid early
degradation by proteolytic enzymes in the upper GI tract or by
proteolytic enzymes in the same formulations. Digestive enzymes
formulations that include trehalase in a time-delayed release form,
should be used along with trehalase combined with antibiotics
(specific for pathogenic microorganism involved or polymicrobial
antibiotics, anti-parasitic, anti-helmintic and anti-protozoa
drugs) and/or natural antimicrobials active against the pathogens
involved. Also, optionally, trehalase can be included in
combinations of digestive enzymes and specific antibiotics and/or
natural antimicrobials for colonic irrigation treatment method in
single-step or multi-step treatment procedures (as described for
biofilm treatment on mucosal linings in the previous
paragraphs).
Dental and Periodontal Diseases
The two groups of bacteria responsible for initiating dental
caries, including Streptococcus mutans and Lactobacillus (known to
possess multiple pathways for biosynthesis of trehalose), have
direct access to high concentrations of orally ingested simple
sugars and other saccharides, as well as those produced by the
action of salivary amylase on ingested carbohydrates
(polysaccharides), that favors the increased synthesis of trehalose
and formation of microbial biofilm. Enzyme trehalase in combination
with antimicrobials (ex. Chlorhexidine Gluconate), and/or with
other saccharidases (ex. Dextranase) can be used for prevention of
dental caries by inhibiting the formation of bacterial biofilms on
the teeth and surrounding tissue surfaces.
Periodontal disease is a classic biofilm-mediated condition that is
refractory to treatment by antimicrobials alone. Applied
treatments, which include combination of trehalase with
antimicrobials (ex. Chlorhexidine Cluconate) and added other
saccharidases (ex. Dextranase), can be both preventive and
curative. Combination of trehalase with antimicrobials (alone or
with other saccharidases) can be used in oral application for
treatment of periodontal diseases and/or during a professional
dental cleaning procedure. Also, the multi-step local treatment,
including the application of trehalase in combination with
antimicrobials (alone or with other saccharidases), followed by the
application of proteolytic, fibrinolytic, and lipolytic enzymes,
and finally by the application of trehalase in combination with
antimicrobials, as disclosed above for treating infectious biofilm
on mucosal linings, can be used as a curative method for
periodontal biofilm-based infections. Since the bacterial biofilm
is the essence of the dental plaque, the use of trehalase in
combination with antimicrobials (alone or with other saccharidases)
in the mouthwash or gel form can diminish the formation of the
dental plaque, and in prolonged use of trehalase in combination
with antimicrobials can gradually degrade and eliminate the
existing bacterial biofilms.
For dental surgery, the use of combination of trehalase with
antimicrobials in prepared formulations can serve as prophylaxis
against biofilm-based infections. Trehalase in combination with
antimicrobials (ex. Chlorhexidine Gluconate) can be used in pre-
and post-operative dental surgery procedure. Additionally, it can
be combined with the other materials commonly used to treat teeth
in endodontics, such as dental cements.
A prophylactic application of trehalase in combination with
antimicrobials (including natural antimicrobials, such as medicinal
plant-derived essential oils or their active compounds) in dental
hygiene includes its use in mouthwashes, toothpastes, dental floss,
and chewing gum. Trehalase in combination with antimicrobials can
be included into conventional non-alcohol-containing mouthwashes
(to avoid alcohol-induced denaturation of the enzyme); such
compositions also typically include menthol, thymol, methyl
salicylate, and eucalyptol. Inclusion of trehalase in combination
with antimicrobials in toothpaste is straightforward, without
chemical interaction with components of conventional toothpaste;
typical toothpaste formulations comprise: abrasive 10-40%,
humectant 20-70%, water 5-30%, binder 1-2%, detergent 1-3%, flavor
1-2%, preservative 0.05-0.5% and therapeutic agent 0.1-0.5%.
Impregnation of dental floss fibers with trehalase in combination
with antimicrobials is analogous to inclusion of flavorings used in
dental floss materials such as silk, polyamide, or Teflon. Finally,
trehalase in combination with antimicrobials (alone or with other
saccharidases) can be included in chewing gum compositions to
prevent the formation of dental plaques and bacterial biofilms, as
well as to treat oral biofilm-based infections in treatment
protocols with antimicrobials.
Mitigation of Ingestion of Excess Trehalose by Susceptible
Individuals
Owing to its unique chemical structure and properties, trehalose
remains stable under low pH conditions, even at elevated
temperatures, and has the ability to protect proteins in a wide
range of temperature changes, including deep freezing, that makes
it an attractive substance for use as a stabilizer and conservant
for various products in medical and food industries. Over the last
two decades, the agro-food industry has introduced the use of
trehalose in many foodstuffs as a food stabilizer, sweetener, and a
moisture retainer, since the high stability of trehalose enables
the original product characteristics to be retained even after heat
processing, freezing, and prolonged storage. Usually, the product
labeling does not indicate the presence or amount of this food
additive. Patients exhibiting biofilm-based infections, especially
those with genetic trehalase enzyme deficiency, can be at increased
risk upon consumption of the dietary trehalose, as the excess of
this sugar can be used by the gut bacteria for local GI tract
biofilm formation. For mitigation of these negative events, enzyme
trehalase can be added in a time-delayed release form to existing
formulations of digestive and systemic enzymes, to avoid negative
consequences upon consumption of excess amount of dietary
trehalose.
Embodiments for the Treatment of Biofilm-Based Infections on
Medical Devices and Medical Equipment
The methods for treatment of biofilm-based infections on medical
devices and medical equipment exposed to bodily fluids and tissues
comprise two categories: preventive and curative. The preventive
methods of the present compositions and methods rely on altering
the device and equipment surfaces by using trehalase in combination
with antimicrobials, whereas curative methods exploit temporary
exposure of such surfaces to treatment compositions of trehalase
with antimicrobials and other compounds in various treatment
protocols.
Preventive Methods
To prevent microbial biofilm growth on medical devices and
equipment surfaces, trehalase in combination with antimicrobials
can be used in coatings (both delayed-release and non-delayed
release) and for immobilization on the surfaces of such devices and
equipment. Simple (non-delayed release) coatings, comprising
trehalase with antimicrobials, can be applied to metal and polymer
surfaces, and fabric materials to provide a brief initial exposure
of biofilm and biofilm-forming pathogens to treatment combinations.
Delayed-release coatings can release trehalase and antimicrobials
into the surrounding environment over time to gradually degrade
biofilm and kill the embedded pathogens, ultimately depleting the
initial amount of coating contained enzyme and antimicrobials. In
contrast to these coatings, when combinations of trehalase with
antimicrobials are immobilized on a surface of medical devices and
equipment, the enzyme can act as a permanent, reusable catalyst,
providing the potential for ongoing degradation of biofilm and
providing antimicrobials with continuous direct access to
biofilm-embedded pathogens.
Treatment coatings, containing trehalase in combination with
antimicrobials, can be applied to non-porous surfaces (metal and
polymer surfaces of medical devices and equipment) and porous
surfaces, such as those of fabric and fabric-based surgical sewing
material, surgical mesh used for hernia repair, surgical wounds,
burns and skin lesions dressing materials. A foremost example is a
method to prevent biofilm formation and growth on prosthetic heart
valves by impregnating the fabric sewing cuff with trehalase in
combination with antimicrobials before attachment of the cuff to
the heart valve assembly; additionally, the heart valve assembly
can be covered with an immobilized coating comprising trehalase in
combination with antimicrobials. Delayed-release coatings that
discharge trehalase in combination with antimicrobials over time
offer the prospect of prophylactic action against the formation of
microbial biofilms on the biofilm-vulnerable surfaces of medical
devices and on temporary and permanent bodily implants. The
delayed-release coatings can include combination of trehalase with
antimicrobials embedded in surface porosity either pre-existing or
specially-created at the surface, surface-attached
microencapsulated trehalase with antimicrobials, and dissolvable
coatings overlaying the combination of trehalase with
antimicrobials on the surface. The methods of adhesion to the
device or implant surfaces, and the mechanisms of time release of
agents of interest are well known in the prior art and can be
modified to exploit the use of trehalase in combination with
antimicrobials in the present compositions and methods.
Trehalase in combination with antimicrobials can be immobilized (as
discussed below in greater detail with respect to curative methods)
on the biofilm-vulnerable surfaces of medical devices and
equipment. The methods of enzyme immobilization on polymer and
metal surfaces is described in detail by Drevon G F ("Enzyme
Immobilization into Polymers and Coatings," PhD Dissertation,
University of Pittsburgh, 2002). Immobilization of trehalase with
antimicrobials on the surface of medical devices and equipment can
be combined with other materials of antimicrobial nature, such as
medical silver and medical copper, and/or with biofilm attachment
preventives like Bacticent.TM. KB. Combination of trehalase with
antimicrobials can be immobilized on a compound that serves as a
support structure, and this support structure compound can be bound
to device surfaces. This method insures that trehalase enzymatic
activity is preserved by avoiding possible direct interaction of
trehalase with the device surfaces. From among the numerous
candidate support structure compounds, a choice can be optimized
with respect to maintaining the enzymatic activity of trehalase
while achieving high binding affinity to the device surfaces.
Combination of trehalase with antimicrobials, as treatment coatings
both delayed-release and non-delayed release, as well as
immobilized trehalase in combination with antimicrobials, can be
used on the interior and exterior surfaces of central venous
catheters and urinary catheters, and on the biofilm-vulnerable
surfaces of endoscopes and implants of various types, including
orthopedic implants.
The surfaces of implantable and bodily-inserted devices are targets
of both the immune response and bacterial colonization, a so-called
"race for the surface" (Gristina A, "Biomedical-centered infection:
microbial adhesion versus tissue integration," Clinical Orthopedics
and Related Research, 2004, No. 427, pp. 4-12.). In the case of the
immune response acting first, the macromolecule adhesion and
general inflammatory action can lead ultimately to the enclosure of
the device surface by a nonvascular fibrous capsule which further
can support bacterial colonization and biofilm formation. If
bacterial colonization occurs before overt immune response, biofilm
can form immediately adjacent to the device surface. Since both the
accumulation of host cells at the device surface and bacterial
colonization of the surface have initial macromolecule adhesion in
common, defeat of such adhesion in vivo is synergistic with use of
trehalase in combination with antimicrobials to impede biofilm
formation.
For this purpose, trehalase in combination with antimicrobials can
be combined with new coatings that offer the promise of deterring
macromolecule adhesion to synthetic surfaces. Among examples are
Semprus Sustain.TM. technology, a polymeric approach to harnessing
water molecules at device surfaces to impede macromolecule
attachment, Optichem.RTM. antifouling coating with microporosity
excluding macromolecule contact with the protected device surface,
and zwitterionic coatings (Brault N D, Gao C, Xue H, Piliarik M,
Homola J, Jiang S, Yu Q, "Ultra-low fouling and functionalizable
zwitterionic coatings grafted onto SiO2 via a biomimetic adhesive
group for sensing and detection in complex media," Biosens
Bioelectron., 2010 Jun. 15, 25(10): 2276-2282.) that suggest the
prospect of defeating protein adhesion through the exploitation of
periodic reversal of polarity in the surface coating.
Delayed-release coatings which include trehalase in combination
with antimicrobials, can be used in concert with
macromolecule-repellant coatings in various modes. For example, the
combination of trehalase with antimicrobials time release sites can
be established with adequate density within the confines of a
macromolecule-repellant coating. Alternatively, disparate coatings
can be interleaved in various geometries both parallel and
perpendicular to the device surface.
Curative Methods
Methods of the present compositions and methods that address
degradation and removal of biofilms and associated pathogens from
surfaces involve various soak (immersion) and rinse protocols.
Solutions of trehalase in combination with antimicrobials and other
compounds, such as other enzymes, chelating agents, and stabilizers
are anticipated. In a preferred embodiment of a soak solution, the
present inventive use of trehalase enzyme to degrade the biofilm
gel matrix can be viewed as an important addition to enzyme
mixtures found in such products as the aforementioned Biorem.
Immersive exposure to trehalase-based soak solutions can be
followed by exposure to various biocidal treatments, as are well
known in the prior art, for elimination of biofilm-forming
pathogens. Rinse and soak solutions containing trehalase should be
maintained at the temperature of maximum enzyme activity. Also,
soak and immersion durations should be made sufficient for
effectiveness.
A preferred method of solution-based treatment comprises the
following multi-step procedure:
1--creating a first treatment solution taken from the group
comprising trehalase with antimicrobials and saccharidases in
aqueous or saline solution,
2--creating a second treatment solution taken from the group
comprising: a) proteolytic enzymes in aqueous or saline solution,
b) fibrinolytic enzymes in aqueous or saline solution, and c)
lipolytic enzymes in aqueous or saline solution,
3--creating a third treatment solution taken from the group
comprising: a) biocides in aqueous or saline solution, b)
antibiotics, specific to the infectious agents present in aqueous
or saline solution, or c) polymicrobial antibiotics in aqueous or
saline solution, d) natural antimicrobials of wide range of action
in aqueous or saline solution,
4--flushing or rinsing the surface under treatment with these
solutions (or immersing such surface in these solutions) in the
sequence given.
The exposure time for the treated surface should be sufficient for
effectiveness, and such solution treatments should take place in a
manner that avoids exposure of trehalase with antimicrobials and
other saccharidases to proteolytic enzymes.
This multi-step procedure can be applied to treatment of central
venous and urinary catheters, endoscopes, contact lenses and lens
cases, dialysis system components, dental unit water lines, and
other medical devices that can be subjected to immersion, rinse, or
fluid injection. In the case of dialysis systems, various surfaces
that contact biological fluids must be disinfected: some surfaces
can be immersed in treatment solutions with the option of
ultrasound-assisted cleaning, other surfaces are not immersible and
simply must be soaked and flushed with treatment solutions. Also,
for dialysis system components and dental unit water line
treatment, the aforementioned third solution additionally can
contain chelating agents and enzyme stabilizers.
An alternative avenue of delivery of trehalase in combination with
antimicrobials involves immobilization of such combination by
attachment to a support structure compound of some kind. In
contrast to immobilization on device surfaces, as discussed above,
the combination of trehalase with antimicrobials can be immobilized
on a support structure compound that is in liquid suspension for
use as a treatment liquid. Such immobilization of trehalase in
combination with antimicrobials can permit its extended presence
and repeated use in catalysis. Additionally, it can increase the
enzyme's catalytic efficiency and thermal stability based on the
specifics of its attachment to the support structure. There are
five general categories of such immobilization: a) adsorption, b)
covalent binding, c) entrapment, d) encapsulation, and e)
cross-linking (Walker J M, Rapley R, and Bickerstaff G F,
"Immobilization of Biocatalysts" in Molecular Biology and
Biotechnology, 4th edition, edited by J. M. Walker and R. Rapley,
RSC Publishing, 2007). All such mechanisms are within the scope of
the present compositions and methods. In the delivery of trehalase
in combination with antimicrobials to biofilm, some immediate
implementations of immobilization are envisioned herein. For
example, trehalase in combination with antimicrobials can be
covalently bound to microspheres, as discussed below, or
encapsulated in liposomes after the fashion of U.S. Pat. No.
7,824,557 (which discloses the use of antimicrobial-containing
liposomes to treat industrial water delivery systems). These
delivery mechanisms can be incorporated by uptake into the biofilm
matrix to provide sustained exposure to trehalase in combination
with antimicrobials.
The feasibility of immobilization of trehalase in combination with
antimicrobials is underscored by examples of trehalase
immobilization for various non-treatment purposes that can be found
in the recent research literature. For analytical purposes,
Bachinski et al. demonstrated the immobilization of trehalase on
aminopropyl glass particles by covalent coupling that allowed the
enzyme to retain its catalytic activity (N. Bachinski, A. S.
Martins, V. M. Paschoalin, A. D. Panek, and C. L. Paiva, "Trehalase
immobilization on aminopropyl glass for analytical use," Biotechnol
Bioeng., 1997 Apr. 5, 54(1): 33-39.). For reactor reuse, trehalase
has been immobilized on chitin as well (A. S. Martinsa, D. N.
Peixotoa, L. M. C. Paivaa, A. D. Paneka and C. L. A. Paivab, "A
simple method for obtaining reusable reactors containing
immobilized trehalase: Characterization of a crude trehalase
preparation immobilized on chitin particles," Enzyme and Microbial
Technology, February 2006, Volume 38, Issues 3-4, Pages 486-492.).
The present compositions and methods include immobilization of
enzyme trehalase in combination with antimicrobials on support
structures that have particular affinity for biofilms. U. S. Patent
Application No. 20060121019 discloses the covalent and non-covalent
attachment of biofilm degrading enzymes to "anchor" molecules that
have an affinity for the biofilm; moieties cited as having a known
affinity for biofilms included Concanavalin A, Wheat Germ
Agglutinin, Other Lectins, Heparin Binding Domains, enzyme
Elastase, Amylose Binding Protein, Ricinus communis agglutinin I,
Dilichos biflorus agglutinin, and Ulex europaeus agglutinin I.
A preferred method of using immobilized trehalase in combination
with antimicrobials in liquid treatment comprises the same
solution-based multi-step procedure outlined above, but using
immobilized trehalase in aqueous or saline suspension. Likewise,
the method is similarly applicable to treatment of the same
categories of medical devices disclosed above.
As mentioned earlier, ensonification of the surface to be treated
can be employed to augment the removal of microbial biofilm
concomitantly with soak and rinse solutions. Apart from the
traditional use of ultrasound for biofilm removal, an additional
modality that is within the scope of the present compositions and
methods is the use of ultrasound to assist enzymatic activity. The
introduction of a low energy, uniform ultrasound field into various
enzyme-containing solutions can greatly improve their effectiveness
by significantly increasing their reaction rate. The process is
tuned so that cavitation does not result in reduction of the enzyme
activity, but rather results in its significant increase.
It has been established that the following specific features of
combined enzyme/ultrasound action are critically important: a) the
effect of cavitation is several hundred times greater in
heterogeneous systems (solid-liquid) than in homogeneous, b) in
water, maximum effects of cavitation occur at .about.50.degree. C.,
which is the optimum temperature for many industrial enzymes, c)
cavitation effects caused by ultrasound greatly enhance the
transport of enzyme macromolecules toward substrate surface and, d)
mechanical impacts, produced by collapse of cavitation bubbles,
provide an important benefit of "opening up" the surface of
substrates to the action of enzymes (Yachmenev V, Condon B, Lambert
A, "Technical Aspects of Use of Ultrasound for Intensification of
Enzymatic Bio-Processing: New Path to "Green Chemistry",
"Proceedings of the International Congress on Acoustics, 2007).
Enzyme reaction rates can be increased by more than an order of
magnitude. In an example of specific enzyme application,
alpha-amylase reaction rates were increased with the use of
ultrasound (Zhang Y, Lin Q, Wei J N, and Zhu H J, "Study on
enzyme-assisted extraction of polysaccharides from Dioscorea
opposite," Zhongguo Zhong Yao Za Zhi. 2008 February, 33(4):
374-377.). For ultrasound-assisted enzyme-based treatment, the
solution-based multi-step treatment previously disclosed, can be
modified to include ensonification of enzyme-containing treatment
solutions and surfaces under treatment.
Embodiments to Address Industrial Biofilms.
There are numerous industrial biofilm treatment approaches that can
be enabled by the use of trehalase enzyme in combination with
antimicrobials. These approaches involve both creation of
appropriate mixtures of trehalase with antimicrobials and other
compounds, and development of methods for delivery of these
mixtures to the sites of biofilm presence.
With respect to treatment mixtures, the combination of trehalase
with antimicrobials can be used alone in aqueous or saline solution
or can be added to compounds that maintain the optimum pH range
(buffer compositions), and metallic ion concentrations that can
maximize the hydrolysis rate of trehalose. Additionally, one or
more combinations of trehalase with antimicrobials can be added to
compositions of dispersants, surfactants, detergents, other
enzymes, and biocides that are delivered to the biofilm in order to
achieve synergistic effects. Also, trehalase in combination with
antimicrobials can be used as a pretreatment step in various
protocols involving other biofilm treatment compounds and/or
methods.
Also, combination of trehalase with antimicrobials can be
immobilized on substrate compounds in liquid suspensions, as
discussed above, for use in industrial treatments, where the
substrate compound may have an affinity for the target of
treatment.
For oil pipelines, an oil-water emulsion containing trehalase in
combination with antimicrobials will provide a dosing opportunity
to treatment of the biofilms within the pipeline. These
emulsion-borne mixtures can include combination of trehalase with
antimicrobials alone, or with additional conventional treatment
compounds such as other biocides, surfactants, detergents, and
dispersants as are well known in the prior art.
A specific treatment embodiment for pipelines involves the
exploitation of annular liquid flow geometries. The annular flow
pattern of two immiscible liquids having very different viscosities
in a horizontal pipe (also known as "core-annular flow") has been
proposed as an attractive means for the pipeline transportation of
heavy oils since the oil tends to occupy the center of the tube,
surrounded by a thin annulus of a lubricant fluid (usually water)
(Bannwar A C, "Modeling aspects of oil-water core-annular flows,"
Journal of Petroleum Science and Engineering Volume 32, Issues 2-4,
29 Dec. 2001, Pages 127-143.). A thin water film can be introduced
between the oil and the pipe wall to act as a lubricant, giving a
pressure gradient reduction. In 8-inch diameter pipes, it has been
shown that, under certain conditions, it is possible to use very
thin water films. For crude oils with viscosities exceeding 2000
mPas, stable operation has proved feasible with as little as 2%
water (Oliemans R V A, Ooms G, Wu H L, Duijvestijn A, "Core-Annular
Oil/Water Flow: The Turbulent-Lubricating-Film Model and
Measurements in a 2-in. Pipe Loop," Middle East Oil Technical
Conference and Exhibition, 11-14 Mar. 1985, Bahrain.). In an
embodiment of the present compositions and methods to address
delivery of trehalase with antimicrobials-containing solutions to
the interior of oil pipelines, the thin water film is replaced by a
trehalase with antimicrobials in aqueous solution. This combination
of trehalase with antimicrobials in aqueous solution will be a
flowing annular layer immediately adjacent to the inner surface of
the pipeline.
Another embodiment of the compositions and methods, addressing
microbial biofilm in the oil pipelines, comprises the exploitation
of magnetic force to deliver trehalase in combination with
antimicrobials to the target treatment sites within pipelines.
Specifically, combination of trehalase with antimicrobials can be
immobilized on a support structure compound that exhibits either
magnetic or preferably ferromagnetic properties. When this
immobilized combination of trehalase with antimicrobials is
released into pipeline flow, a magnetic field exterior to the
pipeline can be used to guide and retain the immobilized
combination of trehalase with antimicrobials in the target vicinity
on the interior of the pipeline. The magnetic field can be
generated by magnetic or electromagnetic means well known in the
prior art. Optimization of this embodiment could include spatial
and temporal variation of the generated magnetic field to achieve
appropriate concentration of trehalase in combination with
antimicrobials at treatment sites in the presence of fluid flow.
Residual magnetism induced in the pipeline wall can be diminished
by methods well known in the prior art.
Dry dock removal of hull biofouling material, including biofilms,
can use aqueous solutions containing trehalase in combination with
antimicrobials in rinse and/or soak protocols. Application of
hydrogel containing trehalase in combination with antimicrobials to
ships' hull is another means of ensuring sustained exposure of the
biofilm for hydrolysis of the trehalose component of the biofilm
matrix and initial elimination of biofilm-embedded microorganisms.
This can be done prior to or at the time of additional biocide
application. Further, the biofilm preventive coatings, containing
combination of trehalase with antimicrobials, can be immobilized on
marine surfaces. The solution-based, multi-step treatment that
includes trehalase in combination with antimicrobials discussed for
medical device treatment can be used in the marine surface
applications, or it can be modified to use gel delivery of
treatment compounds instead of aqueous or saline solutions.
For HVAC systems the solution-based multi-step treatment method
that includes combination of trehalase with antimicrobials and
other compounds, can be used as stated for certain components such
as cooling coils and drain pans, or modified so that treatment
compounds can be fed into HVAC ductwork in the form of
aerosols.
Candidate industrial biocides for use along with trehalase in
combination with antimicrobials include popular industrial biocide
products on the market such as Ultra Kleen.TM. manufactured by
Sterilex Corp., Hunt Valley, Md., the active ingredients of which
comprise:
n-Alkyl(C14 60%, C16 30%, C12 5%, C18 5%) dimethylbenzylammonium
chloride; and
n-Alkyl(C12 68%, C14 32%) dimethylethylbenzylammonium chloride.
Another example is SWG Biocide manufactured by Albermarle Corp.,
Baton Rouge, La., the active ingredients of which comprise sodium
bromosulfamate and sodium chlorosulfamate. Candidates may also be
found among the wider generic categories of industrial biocides
comprising: glutaraldehyde, quaternary ammonium compounds (QACs),
blends of Gut and QACs, Amine salts, Polymeric biguanide,
benzisothiazolone, blend of methyl isothiazolones, and acrolein
(Handbook of Biocide and Preservative Use, Edited by H. W.
Rossmoore, Chapman and Hall, 1995).
For treatment of biofilms associated with food processing, storage,
and transport systems, conventional enzyme treatments can be
augmented with the use of trehalase in combination with
antimicrobials. This can be done in the context of the
solution-based multi-step procedure. In addition, the present
compositions and methods include the use of trehalase in
combination with antimicrobials and other enzymes in one-step
procedure, given that added enzymes are not proteolytic. Also,
ultrasound-assisted enzyme-based cleaning is applicable with the
use of trehalase in combination with antimicrobials.
Biofilms are found in the household environment on many surfaces
including the inside surfaces of plumbing and drainpipes, on the
surfaces of sinks, bathtubs, tiling, shower curtains, shower heads,
cleaning sponges, glassware, toothbrushes, and toilets. Aqueous- or
saline-based solutions containing trehalase in combination with
antimicrobials can be used alone or in proper combination with
other biofilm treatment products tailored to the applicable
surfaces in cleaning procedure. For example, certain compounds used
for plumbing treatment would be inadmissible for treating
toothbrushes. The aforementioned solution-based multi-step
procedure easily can be applied to many household surfaces with the
exception of the internal surfaces of plumbing.
Test Results. There now follows a summary of various in vitro test
results using Trehalase to determine Trehalase potential in
gram-positive and gram-negative bacteria and determine the effect
of Trehalase in combination with the antibiotics ceftazidime (CAZ),
gentamicin (GENT) and tobramycin (TOB) both with grown biofilms,
added at the beginning of experiments, and added to an early
24-hour grown and late 48-hour grown preformed biofilms during a
24-hour exposition. The effect of Trehalase in gram-positive
bacteria (MRSA and MSSA) S. aureus and in gram-negative bacteria P.
aeruginosa is also summarized. Following this summary is a more
detailed explanation of the experimental process with supporting
graphs and charts for the results. The tests were conducted and
summarized by the Drug Discovery and Development Pharmaceutical
Services Company Aptuit, LLC in Verona, Italy.
It is well known that Gram-positive and Gram-negative bacteria use
a ubiquitous multifunctional sugar, i.e., disaccharide trehalose,
as a general stress response metabolite and osmoprotectant, to form
biofilm as a protective cover against harmful environmental
factors, and to preserve integrity of the bacterial cells for
survival in a hazardous environment (including the milieu of the
human body). The results from the testing show that the enzyme
Trehalase, highly specific for degrading disaccharide trehalose
(one substrate--one enzyme), can be effectively used for prevention
and treatment of microbial biofilms, overcome bacterial resistance
to antimicrobials, and increase the effectiveness of existing
treatment modalities. Due to structural differences of
Gram-positive and Gram-negative bacteria, the effect of applied
enzyme Trehalase can have some special features in both types of
bacteria.
Because Trehalose is a structural element in the cell wall of
Gram-positive bacteria, the use of the external enzyme Trehalase
can affect not only biofilm formation, but also can exert certain
effects on the bacterial cell wall and cell membrane, increasing
their permeability to antibiotics, and possibly decreasing
antimicrobial resistance. Gram-negative bacteria has the intrinsic
enzyme Trehalase present in the outer membrane space (acidic
Trehalase) that degrades trehalose released from the cytosol for
recycling, and metabolizes trehalose taken from outside the cell
for intracellular utilization. In Gram-negative bacteria, the
application of external enzyme Trehalase may require more time for
addressing the biofilm formation, and taking longer to exert an
effect on the cell membranes, but still influencing their
permeability to antibiotics and decreasing antimicrobial resistance
in many cases.
The results of in vitro studies performed by Aptuit, LLC
demonstrated some differences in the effect of the enzyme Trehalase
(alone and in combination with antibiotics) on microbial biofilms
produced by Gram-positive and Gram-negative bacteria. The enzyme
tested in the study was an enzyme of mammalian origin, i.e., pig
kidney Trehalase, produced by "Sigma-Aldrich" (USA), which in some
tests was later purified by a dialysis procedure developed by
Aptuit/Verona S.r.l. as will be explained briefly below.
Various parameters that were evaluated in each part of the analysis
include: 1) Biofilm mass formation (% inhibition--by Crystal Violet
staining method); 2) Bacterial cell viability (% inhibition--by
Resazurin assay); and 3) Bacterial cell growth (by Colony
Counting--CFU/ml plating).
One in vitro test was an exploration of the Trehalase potential in
gram-negative bacteria and an exploration of Trehalase Potential in
Gram-Negative Bacteria (P. aeruginosa PAO1--reference strain), and
which showed an effect of Trehalase alone added to the media (TSB)
at the beginning of a 72 hour experiment. Trehalase alone inhibited
biofilm mass formation in a dose-dependent manner. From various
doses (0.023, 0.046, and 0.092 U/well) the highest effect was seen
with 0.092 U/well. The "U" corresponds to the International Units
for the enzyme in each testing well. This dose was further used in
subsequent experiments. Compared to a control, Trehalase alone
inhibited the biofilm mass formation during 72 hours of biofilm
growth, with the most significant inhibition noted at the initial
(4 hour) stage of biofilm development at 83.3%. In an early (24
hour) mature biofilm inhibition was 82.4%, and in fully mature (48
hour) biofilm inhibition was at 91.0%. In an old stage (72 hour of
growth), the biofilm formation was still inhibited by about 50.0%.
It appears that Trehalase alone did not affect the bacterial cell
viability and growth at various stages of the biofilm development
(4 hours, 24 hours, and 48 hours), as confirmed by Resazurin
assessment and bacterial counting (CFU/ml), i.e., colony-forming
unit per milliliter. At the same stages of biofilm development (4
hours, 24 hours, and 48 hours), no effect on biofilm formation and
cell viability was seen with solvent present in the Sigma Trehalase
preparation (Glycerol/water 50/50 solution and 1% Triton-X100) used
in the testing.
A next series of tests explored the effect of Trehalase in
combination with the antibiotics Ceftazidime (CAZ) and Tobramycin
(TOB) in 1/4 and 1/8 sub-MIC (Minimum Inhibitory Concentration)
doses in 24 hour to 48 hour grown biofilms that were added at the
beginning of the experiments. Trehalase alone demonstrated
continuing inhibitory effect on the biofilm mass formation with
longer exposition time (about 65% inhibition in a 48 hour grown
biofilm) with no effect on bacterial cell viability and growth in
both 24 hour and 48 hour grown biofilms. Both antibiotics (CAZ and
TOB) in sub-MIC doses (1/8 and 1/4 MIC), introduced to the media at
the beginning of experiment, triggered the Biofilm Induction
Response (BIR) in 24 hour grown and in 48 hour grown biofilm,
increasing biofilm mass formation by 1.2 to 1.3 (CAZ) and 1.3 to
1.7 fold (TOB) compared to the Control level, i.e., the level of
biofilm mass formed by P. aeruginosa PAO1 in the absence of
antibiotics. When Trehalase was added to antibiotics, it abrogated
the biofilm induction response "BIR" to both antibiotics (CAZ and
TOB) in the 24 hour grown biofilm down to the Control level, with
an additional reduction (about 30%) in the biofilm mass formation
only for the Trehalase/TOB combination. With a longer exposition
time in the 48 hour grown biofilm, Trehalase added to antibiotics,
demonstrated higher effect, abrogating biofilm induction response
"BIR" to both antibiotics down to the Control level, with an
additional reduction of biofilm mass formation by about 65% for the
Trehalase/CAZ combination and by about 70% for the Trehalase/TOB
combination (p<0.001; p<0.001 compared to the Control).
Tobramycin alone (at a dose of 1/4 MIC) inhibited cell viability
(by Resazurin assay) by about 25% in both the 24 hour and 48 hour
grown biofilms. The Tobramycin/Trehalase combination enhanced this
inhibitory effect up to 45% only in the 24 hour grown biofilm.
Ceftazidime alone at a dose of 1/8 and 1/4 MIC inhibited cell
viability (Resazurin assay) by about 25% to 50% only with the
longer exposition, i.e., in the 48 hour grown biofilm. The
CAZ/Trehalase combination did not enhance this effect.
A next series of tests explored the effect of Trehalase alone and
in combination with the antibiotics Ceftazidime (T+CAZ) and
Tobramycin (T+TOB) in 1/4 MIC doses, added to the early (24 hour
grown) preformed biofilm during the 24 hours of further exposition.
When added to the early (24 hour grown) pre-formed biofilm,
Trehalase alone inhibited further biofilm mass formation by about
30% (p=0.08) (during the 24 hour exposition). Both antibiotics (CAZ
and TOB) in sub-MIC doses (1/4 MIC) added to the early (24 hour
grown) preformed biofilm, triggered "BIR", increasing biofilm mass
formation by about 1.6 (CAZ) and about 1.4 (TOB) compared to the
Control level (i.e. the level of biofilm mass formed by P.
aeruginosa PAO1 in the absence of antibiotics). Trehalase in
combination with antibiotics, added to 24 hour preformed biofilm,
abrogated "BIR" to both antibiotics, and additionally slightly
reduced biofilm mass formation by about 10% (T+CAZ) and about 37%
(T+TOB) compared to the Control level. Trehalase alone and in
combination with both antibiotics (T+CAZ and T+TOB) as added to a
24 hour preformed biofilm did not inhibit bacterial cell viability
and growth by Resazurin assay and Colony counting (CFU/ml).
For the Gram-negative pathogen P. aeruginosa PAO1, trehalase alone
added at the beginning of the experiment significantly inhibited
the biofilm mass formation during the 72 hours of biofilm growth,
with the highest inhibition noted at the initial (4 hour) stage of
biofilm development at 83.3%. In the early (24 hour) mature
biofilm, the inhibition was 82.4%. In the fully mature (48 hour)
biofilm, the inhibition was 91.0%. In the old stage of growth (72
hour), the biofilm formation was still inhibited by about 50.0%.
Trehalase alone did not appear to affect bacterial cell viability
and growth at various stages of the biofilm development (4 hour, 24
hour, 48 hour, and 72 hour), as confirmed by Resazurin assay and
Bacterial Cell Counting (CFU/ml). Trehalase in combination with the
antibiotics cephalosporin Ceftazidime and aminoglycoside Tobramycin
in sub-MIC doses (1/8 and 1/4 MIC), was effective in the 24 hour to
48 hour grown biofilms, first by abrogating the Biofilm Induction
Response (BIR) to both antibiotics, and second by continuing
inhibition of biofilm mass formation with longer exposition time,
with the additional reduction of biofilm mass by about 65% (T+CAZ)
and about 70% (T+TOB) (p<0.001; p<0.001 compared to the
Control), which occurred in the 48 hour grown biofilm. Trehalase
slightly potentiated the inhibitory effect of Tobramycin on biofilm
cell viability (by Resasurin assessment) in the 24 hour grown
biofilm by about 45% inhibition for the Trehalase/Tobramycin
combination, compared to about 25% inhibition for the Tobramycin
alone.
These tests show positive results for the use of Trehalase as an
adjuvant to antibiotics for the prevention and treatment of
biofilm-based chronic infections caused by Gram-negative pathogens,
including drug-resistant bacteria. The Biofilm Induction Response
"BIR" to sub-MIC doses of antibiotics is a known phenomenon and is
considered to be the first step in antimicrobial resistance in
Gram-positive and Gram-negative bacteria, and is important in
clinical practice during the "adjustment" period of antibiotic use,
both at the beginning and the end of the treatment course.
In all of these tests, a Trehalase product from Sigma-Alrich was
used as derived from porcine kidney with a buffered aqueous
glycerol solution of greater than or equal to 1.0 units/milligram
protein. An example product can be acquired from Sigma-Alrich
as:
Product Number: T8778
CAS Number: 9025-52-9
MDL: MFCD00132462
TABLE-US-00001 TEST SPECIFICATION Appearance (color) Colorless to
light yellow Appearance (form) Liquid Appearance (turbidity) Clear
to slightly hazy units/mg protein - one unit will convert 1.0
.gtoreq.1.0 micromole of Trehalose to 2.0 micromoles of glucose per
minute at pH 5.7 and at 37.degree. C. (liberated glucose determined
at pH 7.5) alpha-Galactosidase .ltoreq.1% alpha-Glucosidase
.ltoreq.1% .beta.-Glucosidase .ltoreq.5% Invertase .ltoreq.1%
Amylase .ltoreq.2% mg protein/ml (BCA) 0.5-10.0
There now follows greater details about tests that show the effect
of Trehalase in gram-positive bacteria (MRSA and MSSA S. aureus). A
first test showed the effect of Trehalase on initial biofilm
formation by MRSA S. aureus ATCC 25923 (lab strain). Trehalase
alone (at a dose of 0.092 U/well) was added to the media as a
Trypticase soy broth (TSB) at the beginning of the experiment, and
inhibited the initial biofilm mass growth (p<0.001 compared to
the Control), most significantly (about 85% to about 90%) at the
very early stages (3 hour and 7 hour) of biofilm formation, and
about 67% in the early (28 hour) mature biofilm. It inhibited
viability of bacterial cells in the biofilm (about 84%, about 81%,
and about 63% correspondingly by Resazurin assay), and reduced the
bacterial cell growth in the 24 hour biofilm (about 1.5 log by
CFU/ml compared to the Control). Trehalase did not affect bacterial
cell viability and growth in the population of planktonic cells in
the supernatant removed after 24 hours of biofilm growth.
A second test showed the effect of Trehalase in combination with
the antibiotics Ceftazidime (CAZ) and Gentamicin (GENT) in sub-MIC
doses (1/4 and 1/8 MIC) in early (24 hour grown) biofilm formed by
MRSA S. aureus ATCC 25923 [MIC Ceftazidime on S. aureus ATCC
25923=8 .mu.g/mL by CLSI guidelines]; [MIC Gentamicin on S. aureus
ATCC 25923=2 .mu.g/mL (by CLSI guidelines]. When Trehalase (at a
dose 0.092 U/well) in combination with both antibiotics (CAZ and
GENT) in sub-MIC (1/8 and 1/4 MIC) concentrations, was added to the
media (TSB) at the beginning of experiment, it demonstrated a
synergistic effect on all parameters in the study in the 24 hour
grown biofilm.
There was a synergistic effect of the Trehalase in combination with
Ceftazidime. Biofilm mass formation was inhibited by about 48% (T),
by about 15% to 20% (CAZ), and about 60% (T+CAZ). Biofilm cell
viability (by Resazurin assay) was inhibited by about 62% (T), by
about 13% (CAZ in both sub-MIC doses), and by about 88% to 90%
(T+CAZ in both sub-MIC doses). Biofilm cell growth (CFU/mL) was
reduced by about 1.67 log-1.77 log by T+CAZ in both sub-MIC doses
(p<0.001, p<0.001, p<0.001 compared to the Control,
Trehalase alone and Ceftazidime alone correspondingly). There was
no effect on planktonic cells from supernatant removed after 24
hours of incubation was recorded.
There was a synergistic effect of Trehalase in combination with
Gentamicin. The synergistic effect of Trehalase in combination with
Gentamicin (T+GENT) in both sub-MIC doses 1/8 and 1/4 MIC) in 24
hour grown biofilm was significantly higher than in combination
with Ceftazidime, demonstrating. The inhibition of biofilm mass
formation by about 90% to 92% (p<0.001, compared to the Control)
and the full (100%) inhibition of bacterial cell viability (by
Resazurin assay). It also completely abrogated bacterial cell
growth in both populations of the cells by about 3.73 log in
biofilm cells, and by about 3.31 log in planktonic cells from
supernatant removed after 24 hour of incubation (down to the "limit
of detection," i.e., mean--12500 CFU/mL) (p<0.001, p<0.001,
compared to Control).
A series of tests were conducted to show the effect of Trehalase in
combination with the antibiotics Ceftazidime (CAZ) and Gentamicin
(GENT) on early (24 hour grown) and late (48 hour grown) preformed
biofilms formed by MRSA S. aureus ATCC 25923 [Antibiotic
concentrations used in the experiment: Ceftazidime 2 .mu.g/mL=1/4
MIC; Gentamicin--0.5 .mu.g/mL=1/4 MIC]. Trehalase (at a dose 0.092
U/well) in combination with both antibiotics (CAZ and GENT) in
sub-MIC (1/4 MIC) doses, added to the early (24 hour grown) and
late (48 hour grown) preformed biofilms, demonstrated synergistic
effect on all parameters in the study during the next 24 hour of
exposition, with the higher effect on the late preformed biofilm.
The combination of trehalase with a sub-MIC dose (1/4 MIC) of
Gentamicin (GENT) appeared to be the most efficacious.
Trehalase alone (T) and in combination with both antibiotics, added
to the early (24 hour) preformed biofilm, significantly inhibited
further biofilm growth (during 24 hour exposition) by about 48%
(T), by about 56% (T+CAZ), and by about 54% (T+GENT) [p<0.001,
p<0.001, p<0.001, compared to the Control and antibiotics
alone]. The inhibitory effect was higher in the late (48 hour)
preformed biofilm: about 80% (T), about 70% (T+CAZ), and about 83%
(T+GENT) [p<0.001, p<0.001, p<0.001, compared to the
Control and antibiotics alone].
Trehalase alone, added to the early (24 hour) and late (48 hour)
preformed biofilms, inhibited bacterial cells viability (by
Resazurin assay) approximately to the same extent of about 48% (T).
Trehalase in combination with both antibiotics showed a robust
inhibition of cell viability approximately to the same extent in
both early and late preformed biofilms: by about 70% to 80% (T+CAZ)
and about 70% to 76% (T+GENT). Trehalase alone slightly inhibited
biofilm cell growth in the early preformed biofilm (about 0.96
log), and showed higher inhibition in the late preformed biofilm
(about 1.6 log, p<0.05 compared to the Control).
Trehalase in combination with Ceftazidime (T+CAZ) inhibited biofilm
cell growth in both early and late preformed biofilms (p<0.001),
with some higher effect of combination T+CAZ in the early preformed
biofilm (about 1.96 log) than in the late preformed biofilm (about
1.64 log) [p<0.01, compared to control], and by about 1.53 log
[p<0.01] compared with CAZ alone in early preformed biofilm.
Trehalase in combination with Gentamicin (T+GENT) significantly
inhibited biofilm cell growth in both early and late preformed
biofilms (by about 3.06 log and about 3.49 log correspondingly,
p<0.001, p<0.001 compared to the Control). It also
significantly inhibited biofilm cell growth compared to inhibition
by Gentamicin alone, i.e., by about 2.61 log (p<0.01) in the
early preformed biofilm, and by about 3.26 log (p<0.001) in the
late preformed biofilm.
A next series of tests evaluated a purified (dialyzed) Trehalase on
bacterial growth, cell viability and biofilm mass formation in
early biofilm (24 hour grown) and early preformed biofilm
(preformed for 24 hours) produced by S. aureus ATCC 25923
[experimental dialysis of Sigma Trehalase using "Amicon--R
Ultra--15 Centrifugal Filter Devices" (Sigma-Aldrich), method
modified by Aptuit].
In the early (24 hour grown) biofilm, dialyzed Trehalase (at a dose
0.092 U/well) added at the beginning of experiment, inhibited the
biofilm mass formation by about 72% (p<0.001, compared to
Control), with no effect on biofilm cell viability (by Resazurin
assay) and growth (by colony counting--CFU/mL). No inhibitory
effect on biofilm mass formation, biofilm cell viability and growth
was detected with solvent (potassium phosphate buffer) in dialyzed
Trehalase. Dialyzed Trehalase (at a dose of 0.092 U/well) added to
24 hour preformed biofilm, inhibited bacterial cell growth (by
colony counting): T-1.36E+08 CFU/mL (p<0.001, compared to the
Control--1.06E+09 CFU/mL). Potassium phosphate buffer (solvent in
dialyzed Trehalase) did not show any inhibitory effect on bacterial
cell growth in 24 hour preformed biofilm.
These series of tests show that the dialyzed porcine kidney
Trehalase (prepared with potassium phosphate buffer as a solvent)
was confirmed to be at least as active as the Sigma Trehalase
enzyme (prepared with its solvent containing: 50% glycerol/water
solution+1% Triton-X100 and 25 mM potassium phosphate, pH 6.5). The
performed dialysis reduced to the minimum the possibility of the
inhibitory effect of Sigma Trehalase solvent on biofilm formation
and viability of the bacterial cells.
Tests were conducted to show the effect of dialyzed Trehalase on
biofilm formed by Gram-positive Bacteria (lab strains and clinical
isolates: MRSA S. aureus ATCC25923, S. aureus ATCC33591, 3-Belgium,
and VRE/VSE E. faecalis ATCC29212; IH1851165; E. cocco 14) in 24
hour grown biofilm. Due to some negative effect of potassium
phosphate buffer on VRE/VSE E. faecalis ATCC29212, IH851165, E.
cocco 14, the following studies were performed only on MRSA and
MSSA S. aureus laboratory strains and clinical isolates.
Results showed the effect of Trehalase alone and in combination
with the antibiotics Gentamicin and Vancomycin in sub-MIC doses
(1/4 MIC) in the early (24 hour grown) biofilms formed by the lab
strains and clinical isolates of S. aureus, regarded as the good
(high), medium, and poor (low) biofilm producers. All strains had
the same sensitivity (MIC) to Vancomycin, but a different
sensitivity to Gentamicin.
The test showed the effect of Trehalase alone and in combination
with antibiotics Gentamicin and Vancomycin in sub-MIC doses (1/4
MIC) in the early (24 hour grown) biofilm produced by MRSA S.
aureus ATCC25923, which is a lab strain known as a "good (strong)
biofilm producer" (Gentamicin: MIC=0.125 .mu.g/ml; Vancomycin:
MIC=2.0 .mu.g/ml). There is strong reproducibility of the data
generated with S. aureus ATCC25923 (similar activity among
different experiments). In a 24 hour grown biofilm, Trehalase
alone, at a dose of 0.092 UI/well, significantly inhibited biofilm
mass formation by about 90% (p<0.001, compared to Control) and
decreased biofilm cell viability by about 40% (by Resazurin assay).
Both antibiotics Gentamicin (GENT) and Vancomycin (VAN) in sub-MIC
doses (1/4 MIC) triggered the biofilm induction response (BIR),
that resulted in increased biofilm mass by two-fold (GENT) and
1.8-fold (VAN) (p<0.001, p<0.001, compared to the Negative
Control), with no effect on cell viability and growth. Trehalase in
combination with antibiotics, the abrogated biofilm induction
response to both antibiotics and further significantly decreased
biofilm mass formation to the same degree as Trehalase alone did:
by about 90% (for both combinations T+GENT and T+VAN, p<0.001,
p<0.001, compared to the Control), and also inhibited biofilm
cell viability by about 50% (T+GENT) and by about 30% (T+VAN) (by
Resazurin assay).
Tests also showed the effect of Trehalase alone and in combination
with the antibiotics Gentamicin and Vancomycin in sub-MIC doses
(1/4 MIC) in the early (24 hour grown) biofilm produced by MSSA S.
aureus ATCC33591, known as a "medium biofilm producer" (Gentamicin:
MIC=2.0 .mu.g/ml; Vancomycin: MIC=2.0 .mu.g/ml). Trehalase alone,
at a dose 0.092 UI/well, significantly inhibited biofilm mass
formation by about 67% (p<0.001, compared to the Control), with
no effect on biofilm cell viability (by Resazurin assay) and growth
(by colony counting as CFU/mL). Neither Gentamicin (GENT), nor
Vancomycin (VAN) in sub-MICc doses (1/4/MIC) triggered biofilm
induction response. They both even showed a slight reduction in
biofilm mass formation: about 10% (GENT) and about 20% (VAN)
compared to the Control. Trehalase in combination with both
antibiotics, significantly inhibited biofilm mass formation to the
same degree as Trehalase alone did (by about 67% for both
combinations T+GENT and T+VAN, p<0.001, p<0.001, compared to
the Control). Trehalase in combination with Gentamicin (T+GENT)
significantly inhibited biofilm cell viability (about 80% by
Resazurin assay), and significantly reduced bacterial cell growth
(p<0.05, compared to negative Controls, by colony counting as
CFU/mL) in populations of both biofilm cells and planktonic cells
removed from supernatant after 24 hour of biofilm growth. No such
effects were recorded for Trehalase in combination with
Vancomycin.
The tests showed the effect of Trehalase alone and in combination
with antibiotics Gentamicin and Vancomycin in sub-MIC doses (1/4
MIC) on biofilm produced by S. aureus 3-Belgium, clinical isolate,
known as a "poor (low) biofilm producer" (Gentamicin: MIC=32.0
.mu.g/ml; Vancomycin: MIC=2.0 .mu.g/ml). Trehalase alone or in
combination with sub-MIC (1/4 MIC) doses of Gentamicin and
Vancomycin did not induce a decrease in terms of biofilm mass
formation, biofilm cell viability (by Resazurin assay) and
bacterial cell growth (CFU/mL).
Further tests were conducted to explore the MIC (Minimum Inhibitory
Concentration) determination with selected S. aureus strains (MSSA
Oxford and MSSA ATCC 35556, and MRSA ATCC 25923) in the presence
and absence of Trehalase [in vitro assay: Gentamicin, Vancomycin,
Ciprofloxacin--MIC in broth according to CSLI guidelines]. Addition
of Trehalase resulted in significant MIC value reduction with
Gentamicin (up to 16-fold and 33-fold with clinical and reference
isolates). No significant effect on Vancomycin MIC and
Ciprofloxacin MIC was observed. There was testing of Trehalase on
Clinical Isolates (S. aureus strains: Oxford, ATCC35556, and ATCC
25923 as the reference strain) at a time-point: 24 hours of biofilm
growth.
Trehalase alone (at a dose 0.092 UI/well), added to the media at
the beginning of experiment, significantly reduced biofilm mass
formation with all three S. aureus strains tested: about 80%
(ATCC25923), about 90% (ATCC35556), and about 50% (Oxford)
(p<0.001, p<0.001, p<0.05 correspondingly, compared to the
Control). In the presence of Trehalase, the biofilm cell viability
(by Resazurin assay) was inhibited with all tested S. aureus
strains: by about 40% (ATCC25923), about 25% (ATCC35556), and about
60% (Oxford). Biofilm cell growth (by colony counting as CFU/mL)
was significantly reduced (p<0.05 compared to Control) in
biofilms formed by S. aureus ATCC25923 (lab reference strain) and
S. aureus Oxford (clinical isolate). No effect on planktonic cell
growth was recorded. Overall, Trehalase added to the media at the
beginning of experiment with the early (24 hour grown) biofilm,
induced a reduction in biofilm mass formation and inhibition of
viability of all three selected MSSA and MRSA strains at a 24 hour
time-checking point.
Further tests showed the effect of Trehalase, added during the
initial biofilm growth (up to 24 hours) and after 24 hours of
initial biofilm growth (up to 24 hours of further incubation) on a
catheter segment (14-gauge Teflon intravenous catheter): in vitro
assay with S. aureus XEN 29, adapted by Kadurugamuwa et al., 2003).
The bioluminescence signal on catheter was detected with the IVIS
Lumina image system, and bacterial count (both biofilm and
planktonic cells) was evaluated by colony counting (CFU/ml, agar
plating). The addition of Trehalase at the beginning of the initial
24 hour biofilm growth, induced a significant reduction in biofilm
mass formation on a catheter segment (inner and outer surfaces) as
demonstrated by significantly lower intensity of bioluminescence
signal with S. aureus XEN 29+Trehalase 0.092 UI, compared to the
Negative Control (Blank), Positive Control (S. aureus XEN 29), and
Vehicle (S. aureus XEN 29+buffer content 25 mM potassium phosphate,
pH 6.5). Bacterial cell growth (CFU/ml) was significantly reduced
in both biofilm cells and population of planktonic cells
(p<0.05, p<0.05, compared to Positive Control). There was no
effect of Trehalase added after 24 hours of biofilm growth on
catheter segment in terms of bioluminescence signal intensity and
bacterial cell growth (CFU/ml), compared to Positive Control and
Vehicle.
There now follows further details of the various experimental
designs and materials and methods used to conduct many of the tests
explained above. Various charts and tables that reflect the tests
and conclusions drawn from these tests are also explained with
reference to the drawing figures.
A number of the tests explained above evaluated the potential
synergistic effect of a combination between trehalase and three
antibiotics (at sub-MIC concentrations) on bacterial growth,
viability and biofilm mass in early (24 hours) and late (48 hours)
preformed biofilms produced by S. aureus ATCC25923 and P.
aeruginosa PAO1. The experimental design in this test for a first
strain included: Strain: P. aeruginosa PAO1 Temperature:
35.+-.2.degree. C. Incubation Conditions: Static Time Points: 24-48
hours preformed biofilm Treatment: (added to well containing a
suspension of P. aeruginosa PAO1 .about.10.sup.7 CFU/ml) Solvent
Content: 50% glycerol containing 1% Triton.TM. X-100 and 25 mM
potassium phosphate, pH 6.5 Trehalase Content: 0.092 UI Antibiotic
Concentration: Ceftazidime 0.25 ug/ml (1/4 MIC) Ceftazidime 0.25
ug/ml+Trehalase 0.092 UI Tobramycin 0.06 ug/ml (1/4 MIC) Tobramycin
0.06 ug/ml+Trehalase 0.092 UI Parameters Assessed: Biofilm mass
formation (staining with crystal violet 1%) Cells viability
(incubation with resazurin) Cell growth by colony counting
(CFU/mL)
The experimental design in this test for the second strain
included: Strain: S. aureus ATCC25923 Temperature: 35.+-.2.degree.
C. Incubation Conditions: Static Time Points: 24-48 hours preformed
biofilm Treatment: (added to well containing a suspension of S.
aureus ATCC25923 .about.10.sup.7 CFU/ml) Solvent Content: 50%
glycerol containing 1% Triton.TM. X-100 and 25 mM potassium
phosphate, pH 6.5 Trehalase Content: 0.092 UI Antibiotic
Concentration: Ceftazidime 2 ug/ml (1/4 MIC) Ceftazidime 2
ug/ml+Trehalase 0.092 UI Gentamicin 0.5 ug/ml (1/4 MIC) Gentamicin
0.5 ug/ml+Trehalase 0.092 UI Parameters Assessed: Biofilm mass
formation (staining with crystal violet 0.06%) Cells viability
(incubation with resazurin) Cell growth by colony counting
(CFU/mL)
FIGS. 3 and 4 are the tables of the conclusions for these first and
second strains.
Another series of tests evaluated the dialyzed trehalase on
bacterial growth, viability and biofilm mass in early biofilm (24
hours growth) and early preformed biofilm (preformed for 24 hours)
produced by S. aureus ATCC25923 (first strain) and P. aeruginosa
PAO1 (second strain).
The dialysis of trehalase used Amicon.RTM. Ultra-15 centrifugal
filter devices with the following procedure:
1) Added up to 15 mL of sample (1,600 .mu.L of Trehalase
SIGMA+13,400 .mu.L buffer potassium phosphate to the Amicon.RTM.
Ultra filter device;
2) Placed capped filter device into centrifuge rotor (at
4,000.times.g; T=4.degree. C.) for 30 minutes;
3) Removed filtrate (bottom) and exchanged buffer suspension
(filter device);
4) Repeated steps 2 and 3 twice;
5) As a last step recovered 250 .mu.L of ultrafiltrate Trehalase
and re-suspended it with 1,350 .mu.L of buffered suspension to
obtain 1,600 .mu.L of dialyzed trehalase; and
6) Read spectrum of bottom filtrate.
As noted before, the dialysis procedure of Sigma Trehalase was
introduced by using Amicon Ultra-15 Cenrifugal Filter Device. As a
result of this procedure, 1,600 .mu.L of dialyzed trehalase
containing 5 UI (International Units of activity) was in buffer
solution. This activity was the same as in Sigma Trehalase (1,600
.mu.L, containing solvent) before the dialysis. The preparation of
a Trehalase dose of 0.092 UI is used as 0.092 UI/well. The volume
of each well was 200 .mu.L and included 100 .mu.L Dialyzed
Trehalase (or Trehalase Sigma) and 100 .mu.L of bacterial
suspension of P. aeruginosa or S. aureus. To prepare a total volume
100 .mu.L of Trehalase (representing 0.092 UI/well), the test used
40 .mu.L of Trehalase (5 UI activity)+60 .mu.L of TSBG (media), so
that the activity of Trehaalase was 0.092 UI/100 .mu.L, i.e. 0.92
UI/mL, close to 1 UI/mL (1 mL=1000 .mu.L).
In these tests, dialyzed (purified) trehalase in a dose 0.92 UI/mL
of media, inhibited biofilm formation by S. aureus by 72% in 24
hr-grown biofilm. Sigma Trehalase is sold as a liquid solution of
Trehalase (as a proteinous substance in a solvent, but without the
amount of protein by weight), only having activity in UIs. In the
original tests, the concentration is: = or >0.4 units/mg. One
unit will convert 1.0 .mu.mole of trehalose to 2 moles of glucose
per min at pH 5.7 at 37 A.degree. C. (liberated glucose determined
at pH 7.5). This concentration as = or >0.4 units/mg may
represent Trehalase enzyme activity as = or >0.4 units in 1 mg
of protein (i.e. dry weight of enzyme itself). The original assay:
2.0-6.0 mg/ml protein basis (BCA) in a buffered aqueous glycerol
solution. Foreign activity (including a-galactosidase, invertase,
a- and .beta.-glucosidase, and amylase) was < or =1%, confirming
that this enzyme is highly specific only for degradation of
trehalose. The article by Reiko Ishihara et. al., "Molecular
cloning, sequencing and expression of cDNA encoding human
trehalase", Gene 202 (1997) 69-74), has the specific activity of
human trehalase expressed in E. coli, introduced in units/mg of
protein (i.e. protein from lysed bacterial cells).
The experimental design for the first strain on the early biofilm
included: Strain: S. aureus ATCC25923 Temperature: 35.+-.2.degree.
C. Incubation Conditions: Static Time Points: 24 hours Treatment:
(added to well containing a suspension of S. aureus ATCC25923
.about.10.sup.7 CFU/ml) Solvent 1: 50% glycerol containing 1%
Triton.TM. X-100 and 25 mM potassium phosphate, pH 6.5 Solvent 2:
25 mM potassium phosphate, pH 6.5 SIGMA Trehalase: 0.092 UI
Dialyzed Trehalase: 0.092 UI Parameters Assessed: Biofilm mass
formation (staining with crystal violet 0.06%) Cells viability
(incubation with resazurin) Cell growth by colony counting
(CFU/mL)
The experimental design for the first strain on the preformed
biofilm included: Strain: S. aureus ATCC25923 Temperature:
35.+-.2.degree. C. Incubation Conditions: Static Time Points:
Preformed biofilm for 24 hours, 24 hours of further growth
Treatment: (added to well containing a suspension of S. aureus
ATCC25923 .about.10.sup.7 CFU/ml) Solvent 1: 50% glycerol
containing 1% Triton.TM. X-100 and 25 mM potassium phosphate, pH
6.5 Solvent 2: 25 mM potassium phosphate, pH 6.5 SIGMA Trehalase:
0.092 UI Dialyzed Trehalase: 0.092 UI Parameters Assessed: Biofilm
mass formation (staining with crystal violet 0.06%) Cells viability
(incubation with resazurin) Cell growth by colony counting
(CFU/mL)
The experimental design for the second strain on the early biofilm
included: Strain: P. aeruginosa PAO1 Temperature: 35.+-.2.degree.
C. Incubation Conditions: Static Time Points: 24 hours Treatment:
(added to well containing a suspension of P. aeruginosa PAO1
.about.10.sup.7 CFU/ml) Solvent 1: 50% glycerol containing 1%
Triton.TM. X-100 and 25 mM potassium phosphate, pH 6.5 Solvent 2:
25 mM potassium phosphate, pH 6.5 SIGMA Trehalase: 0.092 UI
Dialyzed Trehalase: 0.092 UI Parameters Assessed: Biofilm mass
formation (staining with crystal violet 0.06%) Cells viability
(incubation with resazurin) Cell growth by colony counting
(CFU/mL)
The experimental design for the second strain on the preformed
biofilm included: Strain: P. aeruginosa PAO1 Temperature:
35.+-.2.degree. C. Incubation Conditions: Static Time Points:
Preformed biofilm for 24 hours, 24 hours of further growth
Treatment: (added to well containing a suspension of P. aeruginosa
PAO1 .about.10.sup.7 CFU/ml) Solvent 1: 50% glycerol containing 1%
Triton.TM. X-100 and 25 mM potassium phosphate, pH 6.5 Solvent 2:
25 mM potassium phosphate, pH 6.5 SIGMA Trehalase: 0.092 UI
Dialyzed Trehalase: 0.092 UI Parameters Assessed: Biofilm mass
formation (staining with crystal violet 0.06%) Cells viability
(incubation with resazurin) Cell growth by colony counting
(CFU/mL)
FIGS. 5 and 6 are summary tables for the test results of the
biofilm produced by S. aureus ATCC25923 (FIG. 5) and the biofilm
produced by P. aeruginosa PAO1 (FIG. 6).
Tests were also conducted to explore the dialyzed trehalase
potential in gram-positive bacteria (reference strains and clinical
isolates). The experiment included:
Step 1: The MIC (Minimum Inhibitory Concentration) activity
determination in S. aureus ATCC25923, ATCC33591, and XEN29 in
combination with Trehalase; based on the data, ATCC33591 and XEN29
strains were replaced with ATCC35556 and Oxford strains.
Step 2: Evaluate the effect of Trehalase on biofilm formation by
different clinical isolates:
Phase 1: Screening of clinical isolates for biofilm formation;
Phase 2: Testing Trehalase on selected clinical isolates.
Step 3: Verify the effect of Trehalase on biofilm growth on Teflon
catheter surface:
Phase 1: Effect of Trehalase added during biofilm growth (up to 24
hours);
Phase 2: Effect of Trehalase (added after 24 hours of initial
biofilm growth) during the next 24 hours from addition).
The MIC determination with selected S. aureus strains proceeded as
a first step. The materials and methods included:
S. aureus strains: Oxford; ATCC35556 and ATCC25923 (as reference
strain)
In vitro assay: MIC in broth (according to CLSI guidelines)
Treatment group (for each strain): 1) Negative control: blank 2)
Positive control: bacterial growth 3) Trehalase:
bacteria+/-trehalase 0.092 UI 4) Gentamicin: compound+/-trehalase
0.092 UI 5) Vancomycin: compound+/-trehalase 0.092 UI 6)
Ciprofloxacin: compound+/-trehalase 0.092 UI
Antibiotic concentration: two-fold series dilution from 16 to
0.0078 .mu.g/mL
Time-point: 24 hours
Incubation: T 37.degree. C.; static condition
The results for the MIC determination with selected S. aureus
strains are shown in FIG. 7. The addition of Trehalase resulted in
the MIC value reduction with Gentamicin (up to 16-fold and 33-fold
with clinical and reference isolates). No significant effect on
Vancomycin and Ciprofloxacin was observed.
The screening of clinical isolates proceeded as a second step
(phase 1) and included the following materials and methods:
S. aureus strains:
Oxford; PK2; 3226; IH-1018129; ATCCBAA1556; ATCC49230;
ATCC35556
S. aureus ATCC25923 as the reference strain
Medium: TSB enriched with 1% glucose (TSBG)
In vitro assay format: 96 well--MTP (Microtiter Plate)
Samples: 4 samples for each bacterial strain
Time-point: 24 hours of biofilm growth
Incubation: T 37.degree. C.; static condition
Read-out: Biomass (0.1% safranin staining)
The results of the screening of clinical isolates are shown in FIG.
8. The S. aureus Oxford and ATCC35556 as the strains were selected
for the next steps (both MSSA).
In the next phase (phase 2), the testing of the Trehalase on
clinical isolates included the following materials and methods:
S. aureus strains: Oxford; ATCC35556 and ATCC25923 as the reference
strain
Medium: TSB enriched with 1% glucose (TSBG)
In vitro assay format: 96 well--MTP
Treatment Groups (treatment added during the growth of biofilm):
Negative control: blank Positive control: S. aureus strains
Vehicle: S. aureus strains+buffer content 25 mM potassium
phosphate, pH 6.5 Trehalase: S. aureus strains+Trehalase 0.092
UI
Samples: 4 samples for each bacterial strain
Time-point: 24 hours of biofilm growth
Incubation: T 37.degree. C.; static condition
Read-out: Biomass (crystal violet), cell viability (resazurin),
planktonic and biofilm bacterial count (agar plating)
The results of the Trehalase testing on clinical isolates are shown
in FIGS. 9A, 9B and 9C and FIGS. 10A, 10B and 10C. Overall, the
addition of Trehalase during the 24 hours biofilm growth induced a
reduction in biomass and viability of selected MSSA strains.
The next step (Step 3) of experiment explored the effect of
Trehalase during the 24-hr initial biofilm growth (added at the
start of experiment) and after preliminary (24-hr) biofilm growth
on catheters. Bacterial biofilm was developed on the catheter
segments (14-gauge Teflon intravenous catheter) by incubating
individual segments into tubes containing a S. aureus Xen 29
suspension in the exponential phase of growth. The bioluminescence
signal on catheter segments was detected with the IVIS Lumina image
system, and bacterial count CFU/ml for biofilm cells and planktonic
cells was evaluated (agar plating). The Step 3--phase 1 explored
the effect of Trehalase added during initial biofilm growth (up to
24 hours) and included the following materials and methods:
Bacterial strain: S. aureus XEN29 (suspension .about.10.sup.7
CFU/ml)
Medium: TSB enriched with 1% glucose (TSBG)
In vitro assay: Biofilm formation on catheter (adapted by
Kadurugamuwa et al., 2003)
Samples: 3-4 samples for each treatment condition
Treatment: Negative control: blank (catheter without bacterial
suspension) Positive control: control infection S. aureus XEN29
(catheter with bacterial suspension) Vehicle: S. aureus
XEN29+buffer content 25 mM potassium phosphate, pH 6.5 Trehalase:
S. aureus XEN29+Trehalase 0.092 UI
Time-point: Treatment added during the incubation--biofilm growth
for 24 hours
Incubation: T 37.degree. C.; static condition
Read-out: Bioluminescence signal and biofilm and planktonic cells
bacterial count CFU/ml (agar plating)
The results for exploring the effect of Trehalase added during
initial biofilm growth (up to 24 hours) on catheter segments are
shown in FIGS. 11A, 11B and 11C. As a second phase, the effect of
Trehalase (added after preliminary 24-hr biofilm growth) during the
following 24 hours of biofilm growth on catheter segments was
explored and used the following materials and methods:
Bacterial strain: S. aureus XEN29 (suspension .about.10.sup.7
CFU/ml)
Medium: TSB enriched with 1% glucose (TSBG)
In vitro assay: Biofilm formation on catheter (adapted by
Kadurugamuwa et al., 2003)
Samples: 3-4 samples for each treatment condition
Treatment: Negative control: blank (catheter without bacterial
suspension) Positive control: control infection S. aureus XEN29
(catheter with bacterial suspension) Vehicle: S. aureus
XEN29+buffer content 25 mM potassium phosphate, pH 6.5 Trehalase:
S. aureus XEN29+Trehalase 0.092 UI
Time-point: Treatment added after 24 hours of initial bacterial
incubation--biofilm growth for further 24 hours
Incubation: T 37.degree. C.; static condition
Read-out: Bioluminescence signal and biofilm and planktonic cells
counting (CFU/ml--agar plating)
FIGS. 12A, 12B and 12C show the results from the testing of
Trehalase added after 24 hours biofilm growth (up to 24 hours). The
addition of Trehalase at the beginning of experiment (during the
initial 24 hours of incubation) induced a significant reduction in
biofilm mass and biofilm cells and planktonic cells growth of S.
aureus XEN29 on the catheter. There was not as much effect of the
Trehalase when added after 24 hours biofilm growth in terms of
bioluminescence signal and CFU.
These tests prompted further study and analysis of the use of
Trehalase with other antimicrobials and ingredients of potential
over-the-counter (OTC) products besides the listed antibiotics. For
example, further analysis and study determined that Trehalase may
be combined with silver in an effective amount so that the
Trehalase operates to break down the biofilm and help the host
cells, for example, macrophages, to obtain access to kill the
bacteria and eliminate them from the human body together with any
accumulated silver. The breakdown of the biofilm with the added
Trehalase allows the silver to operate in a more effective manner
against infectious pathogens with an absence of toxicity and
immunogenicity for humans and animals. It is known that silver
exhibits low toxicity in the human body and there is minimal risk
due to clinical exposure by inhalation, ingestion and dermal
application. It is possible to use colloidal silver preparations
and silver sulphadiazine or the more popular silver nitrate. This
is especially effective when it can be used as coatings for
indwelling catheters and cardiac devices in combination with the
Trehalase. With the emergence of antibiotic-resistant strains such
as CA-MRSA and HA-MRSA as flesh-eating bacteria, there is renewed
interest in using silver as an antibacterial agent and it has been
determined from the study that the Trehalase may make the silver
even more efficient and effective as an antibacterial agent with
the breakdown of biofilm.
The silver source releases silver ions that are effective as an
antimicrobial and based on the conformational changes in
trans-membrane proteins, the Trehalase helps break down more of the
biofilm and parts of the matrix and may ease entry of silver ions
into the cell. Different compounds may be used both in medical
devices, textiles and in ointments for dermatological applications,
including metallic silver that includes nanocrystalline forms in
silver coatings. Phosphate silver compounds have moderate ionizing
capacity while the silver nitrate has very high ionizing capacity
and is commonly used with some dermatological carriers. Silver
chloride has low ionizing capability while silver sulfate has a
moderate ionizing capability. The sulphadiazine complexes have high
ionizing capacity and colloidal silver preparations have moderate
to high ionizing capacity. All these compounds may be used in
combination with the Trehalase in effective amounts, including
ointments, sprays and other applications. Nanochemistry techniques
can be used to produce micro-fine silver particles of less than 20
nm diameter with increased solubility and release of silver ions at
about 70 to 100 ppm. The disruption produced by the Trehalase
facilitates the silver ion interaction. Typically, the ionization
of silver metal is proportional to the surface area of a particle
that is exposed.
It is possible to use a sustained silver release dressing on a
tissue dressing in combination with Trehalase. The silver nitrate
should not exceed 1% in contact with living tissue and at 0.5% has
been effective in inhibiting P. aeruginosa. In combination with the
Trehalase, it is even more effective. Silver sulphadiazine combines
the antibiotic properties of silver with sulphonamide and avoids
the disadvantage related to silver nitrate and may be used at 1% in
a cream base in combination with an effective amount of Trehalase.
It may be combined with polyethylene glycol plus
poly-2-hydroxyethyl methacrylate, liposomes, poly-L-leucine or even
cadaver skin. It may be combined with a lipocolloid formulation
such as 1% to 5% silver sulphadiazine at about 0.25 to 0.3% of
Trehalase in a non-limiting example and up to 1% or 2%. In a
preferred embodiment, it should include 0.5 to 10.0 mg protein per
ml based on the BCA (bicinchoninic acid) and dry weight of the
enzyme, and with 0.4 units in 1 mg of protein optimal and with the
range of 0.2 to 1.0.
In terms of the dressing, the silver could be about less than 10 mg
per 100 cm.sup.2 to more than 100 mg per 100 cm.sup.2 with an
effective amount of Trehalase of a similar 0.5 mg to 10 mg/cm.sup.2
and with a proper enzyme unit activity. This amount may vary
depending on the type of Trehalase. Silver ions may be incorporated
into a substance and released slowly with time as with silver
sulfadiazine or may come from ionizing the surface of a solid piece
of silver as with silver nanoparticles. Although the action is not
positive, it is observed that some cells exposed to silver
(Ag.sup.+) ions may have activated a stress response that lead to
the condensation of DNA in the center of the cell and there is some
cell membrane detachment from the cell wall, cell wall damage and
electron dense granules outside and sometimes inside the cell.
Thus, the silver may help inactivate proteins by binding the
sulfur-containing compounds. Further research and development has
observed that the silver in combination with the Trehalase may
disrupt the biofilm and provide enhanced structural change to help
inactivate proteins even more. This is found to be effective with
gram-negative bacteria that may sustain more structural damage than
gram-positive bacteria such as E. Coli for the gram-negative
bacteria as compared to the gram-positive S. Aureus. The silver may
lead to cell shrinkage and dehydration so that in combination with
the Trehalase it is even more effective. This makes sense since
gram-positive bacteria have a charge of peptidoglycan molecules in
the bacterial cell wall and peptidoglycan is negatively charged.
Silver ions are positively charged and more silver may get trapped
by peptidoglycan. With the enhancement and effectiveness from the
added Trehalase degrading the biofilm, this mechanism may trap even
more of the silver.
It is known that silver nitrate usually releases its silver ions
immediately into solution while silver sulfadiazine gradually
releases the majority of its silver ions over an extended period of
time, and thus, may provide a more steady supply of silver ions
that would be more effective perhaps in a wound cream or bandage.
It is possible to use silver salts and other silver components with
silver nanoparticles. The nanoparticles may be spherical or
rod-shaped and triangular. Different sizes may be used such as 1
ug, 12.5 ug, 50 ug or 100 ug. It is possible that the silver
nanoparticles or other silver may be applied to wound dressings and
endrotracheal tubes as a coating, including surgical masks. It has
been found that cotton fibers are more desirable in some cases.
The amounts of Trehalase can vary but for one possible composition,
silver nitrate at 55 ppm may be used in a wound gel and may vary
from 50 to 60 ppm in one example and 40 to 60 ppm in yet another
example. The gel may include a water-based product with glycerin,
carbomer, sodium chloride, silver nitrate and triethanolamine or
any or their equivalents. An equivalent amount of Trehalase may be
added corresponding to in one example to 0.5 to 10 mg/ml and in yet
another example, 0.2 to 0.6 units per mg of protein and for a
specific amount, and 0.4 units in 1 mg of protein. The ppm
equivalence could be similar to the silver in yet another example.
The Trehalase could be 0.5 to 10.0 mg protein/ml. It has also been
determined that the silver-killed bacteria may increase the
antibacterial activity by the "zombies" effect as noted in the
article "Antibacterial Activity of Silver-Killed Bacteria: The
"Zombie" Effect," Scientific Reports, 2015, the disclosure which is
hereby incorporated by reference in its entirety.
It is possible to use an antimicrobial composition such as
disclosed in U.S. Pat. No. 8,568,711, the disclosure which is
hereby incorporated by reference in its entirety, and which
includes the silver ion and 80% water as a solvent including a
hydrophilic polymer where the silver content has the addition of
the Trehalase also ranging from 0.0001 to 0.01 milliliter. It
should be understood that the activity units are to be acceptable
units of USP per milligram.
In one example as a solution, it is possible to use 0.5 to 10 mg/mL
on a protein basis for the Trehalase and yet another example in an
assay 2.0 to 6.0 mg/ml or about 0.4 units/mg. One unit is defined
as the amount of enzyme that may convert 1.0 umol of Trehalose to
2.0 umols of glucose per minute at a pH of about 5.7 at 37 Degrees
C. (the liberated glucose is determined at pH 7.5). Its physical
form may be contained in 50% glycerol containing 1% Triton X-100
and 25 mM potassium phosphate at a pH of about 6.5. The molecular
weight varies but could be about 80,000 Daltons and can range from
70,000 to 85,000 Daltons depending on the origin of the Trehalase
in an aspect. One technique that can be used to prepare the
Trehalase is described in an article from Yoneyama entitled,
"Purification and Properties of Detergent-Solubilized Pig Kidney
Trehalase," Archives of Biochemistry and Biophysics (1987), the
disclosure which is hereby incorporated by reference in its
entirety.
It is possible that Trehalase as alpha, alpha-trehalase, EC
3.2.1.28 could be solubilized from the brush border membrane of pig
kidney cortex by Triton X-100 and sodium deoxycholate in the
presence of inhibitors of proteolytic enzymes. In this example, the
kidney enzyme can be purified 3060-fold using gel filtration, ion
exchange chromatography, Con A-Sepharose chromatography,
phenyl-Sepharose CL-4B hydrophobic interaction chromatography.
Contaminant proteins can be absorbed as described in the article
with 99% or greater purity based on amino-terminal amino acid
analysis. This purified enzyme in this example had a specific
activity of 278 units/mg protein, a molecular weight of about
80,000 on sodium dodecyl sulfate-polyacrylamide gel
electrophoresis. It is a glycoprotein and contained 2 mol of
glucosamine per mole of trehalase. The apparent Km for this
trehalase was calculated to be 2.1 mM. This type of produced kidney
trehalase was highly specific for trehalose and exhibited an
optimal pH of 5.9, and an isoelectric point is between about pH 4.7
and 4.4. Other details may be found in the incorporated by
reference article.
The Trehalase may also be combined with chlorhexidine gluconate or
an equivalent both as an antiseptic skin cleanser and as in an oral
rinse. In one example, the chlorhexidine gluconate is about 0.12%
and in an oral rinse and may be used 3 to 4 times daily. The
Trehalase may be added in units as noted before and in a percentage
if used for labeling at about 0.1% to 0.3% and in yet another
example, 0.1% to 0.5% and up to 1% to 2% in yet another example. It
has been found that a small amount is effective to aid in an oral
rinse for potential applications and to help in breaking down some
biofilm to make the chlorhexidine gluconate more effective. As an
antiseptic skin cleanser, great amounts of the Trehalase may be
used and the higher end range closer to 50 to 100 mg protein/ml and
up to 1% to 7% and in another example about 4% in combination with
4% solution chlorhexidine gluconate. This use of the oral cleaner
and skin cleanser is excellent for the extremely-drug-resistant
(XDR) strains of Klebsiella pneumoniae that may have reduced
susceptibility to chlorhexidine and with the added Trehalase,
allows the chlorhexidine to be more effective since the added
Trehalase helps break apart the biofilm to allow more effective
chlorhexidine.
It is possible to use another decolonization measure such as
mupirocin to prevent Staphylococcus aureus skin and soft tissue
infections (SSTI). It is also possible to use it on the
Acinetobacter baylyi adp1. It may be used in a water-based solution
in one example and for a skin cleanser could be mixed with an
alcohol-based solution of 70% or a water-based solution that may
include other components such as some glycol. The incorporation of
about 2 weight percent and 3 weight percent chlorhexidine in
combination with a 1 to 4 weight percent Trehalase at an effective
enzyme activity is believed to be effective at that range for the
antiseptic skin cleanser. Concentrations may be about or greater
than 1 .mu.g/ml and in some applications, concentrations up to 10
or even greater than 73 .mu.g/ml in some cases as a wipe with
Trehalase.
In one example, it is possible to use 4% w/w for chlorhexidine
gluconate and alcohol at 4% w/w in water with a similar preparation
of the Trehalase. Another possible application is with denture
cleaning tablets that includes a number of components such as a
mild bleach, for example, dilute sodium hypochlorite and may
include other ingredients such as sodium bicarbonate to alkalize
the water and may include citric acid to help remove stains and
sodium perborate, sodium polyphosphate, and potassium
monopersulfate as a cleaning and bleaching agent and EDTA. As a
percent w/w, it may be added to about 1% to 5% for the Trehalase.
It can be added but at an effective 0.5 to 10 mg protein per ml,
and about 0.2 to 1.0 units in a milligram (mg) of protein based on
a dry weight basis, and in an example, 0.4 units/mg. An example of
some the components that may be included with Trehalase include the
following:
TABLE-US-00002 Chemical Name Proportion (% w/w) Potassium
peroxymonosulfate sulfate <10 Sodium carbonate peroxide
10-<30 Sodium carbonate 10-<30 Citric acid 10-<30 Malic
acid <10 Other ingredients classified as not hazardous to 100
according to NOHSC
It is possible to include Trehalase in mouthwashes, toothpastes,
and cement material for dental applications where the Trehalase is
combined with chlorhexidine (as the antimicrobial), or with
essential oils (or their active fractions). For example, some
mouthwashes may include Chlorhexidine Digluconate at 0.06% w/v and
Sodium Fluoride (250 ppm fluoride) and include the added Trehalase
in an amount equivalent to the optimal doses similar to that added
to other compositions and treatments discussed above. Another
example is a mouthwash that may contain as active ingredients
Eucalyptol at 0.092%; Menthol at 0.042%; Thymol at 0.064%, (these
are active antimicrobial fractions of essential oils) and
Methylsalicylate at 0.060%. These components may be labeled
together on the bottle as Antiplaque/antigingivitis. Another type
of mouthwash contains as active ingredient Sodium fluoride 0.02%
(0.01% w/v fluoride ion) along with all ingredients from the first
type (but without their doses, just mentioned as inactive
ingredients). Trehalase can be added to all three mouthwashes in
amount equivalent to the "optimal" doses noted above. Trehalase may
be added to a toothpaste, for example, a type having an active
ingredient as Sodium fluoride at 0.310% w/w (1400 ppm fluoride), or
another toothpaste that has active ingredients such as Sodium
fluoride at 0.24% (0.15% w/v fluoride ion) and Triclosan at 0.30%,
as antigingivitis, i.e., antimicrobial.
As noted before, silver and the Trehalase is effective and it has
also been found that copper is effective. Hospital environments may
act as a reservoir for biofilm-forming pathogens that cause
healthcare-associated infections (HCAIs), so that approaches to
reducing environmental microbial contamination in addition to
cleaning, attracted attention over the last decade. Copper is well
recognized as a powerful antimicrobial with rapid broad spectrum
efficacy against bacteria, viruses and fungi. In a novel cross-over
study in an acute medical ward, a toilet seat, a set of tap handles
and a ward entrance door push plate, each containing copper, were
sampled for the presence of microorganisms and compared to
equivalent standard, non-copper-containing items on the same ward,
demonstrating a statistically significant decrease of microbial
contamination on copper-covered surfaces (A. L. Casey, D. Adams, T.
J. Karpanen, et. al. "Role of Copper in reducing hospital
environment contamination", The Journal of Hospital Infection, Vol.
74, Issue 1, pages 72-77 (January 2010).
Extensive scientific information on the role of Antimicrobial
Copper in reducing transmission of infection and spread of
antimicrobial resistance is introduced in "Scientific References"
(http://www.antimicrobialcopper. or/uk/antimicrobial-resistance) in
multiple laboratory tests carried out under typical indoor
conditions, it was demonstrated that "antimicrobial copper" was
effective against many pathogens, including those with
drug-resistance: MRSA S. aureus, MDR Tubercle bacillus, MDR
Acinetobacter baumannii, Vancomycin-resistant enterococcus (VRE),
Carbapenem-resistant Enterobacteriaceae (CRE), ESBL-producing
Klebsiella pneumoniae, ESBL-producing E. coli.
This efficacy translates into the clinical environment, as
demonstrated in a multi-center ICU trial in the US (M G Schmidt, H
H Attaway, P A Sharpe, et. al., "Sustained Reduction of Microbial
Burden on Common Hospital Surfaces through Introduction of Copper",
Journal of Clinical Microbiology 2012, Vol. 50 No. 7 2217-2223).
Six near-patient surfaces were upgraded to copper, and sampling was
undertaken weekly over a period of 23 months. Over the intervention
period, the combined MRSA and VRE burdens were 96.8% lower on
copper surfaces than on comparable plastic, wood, metal, and
painted surfaces and were 98.8% lower on the bed rails, the most
heavily burdened object. In this US trial, the bioburden reduction
was associated with a 58% reduction in infections (CD Salgado, K A
Sepkowitz, J F John, J R Cantey, H H Attaway, K D Freeman, M G
Schmidt, "Copper Surfaces Reduce the Rate of Healthcare-Acquired
Infections in the Intensive Care Unit", Infection Control and
Hospital Epidemiology 2013, 34(5), 479-486.).
The enzyme Trehalase can be immobilized on the copper-containing
surfaces to prevent microbial biofilm formation by microorganisms
accumulated on such surfaces during the time periods between the
cleaning procedures (frequency of cleaning such surfaces is usually
introduced in guidelines for maintenance of such surfaces). The
enzyme Trehalase can also be included in disinfecting solutions for
such cleaning procedures (as described in previous embodiments for
prevention and treatment of biofilm-based infections on medical
devices and medical equipment surfaces).
Waterborne infections contribute to transmission of infections in
hospital settings, in particular when used in medical equipment
(surgical instruments, endoscope washer-disinfectors, nebulizers,
endrotracheal tubes) disinfection procedures. For example, surgical
instruments may have high post-cleaning levels of carbohydrates (up
to 352 .mu.g/cm.sup.2) and endotoxin (up to 25 373 EU/cm.sup.2),
suggesting unrecognized issues with the quality of water used for
the final rinse, and showing the necessity to monitor the water
quality used in instrument washers (M. J. Alfa, N Olson, A
Al-Fadhaly, "Cleaning efficacy of medical device washer in North
American healthcare facilities", The Journal of Hospital Infection,
Vol. 74, Issue 2, Pages 168-177 (February 2010). The development of
microbial biofilm in washer-disinfectors, the type of biofilms and
the nature of the bacteria within them, along with increasing
antimicrobial resistance in those pathogens is of a major concern
(W. G. MacKay, A. T. Leanord, C. L. Williams, "Water, water
everywhere nor any of a sterile drop to rinse your endoscope", The
Journal of Hospital Infection, Vol. 51, Issue 4, pages 256-261
(August 2002). Growing resistance of HCAIs to antimicrobials and
biocides is on the rise. Various biocides show a wide range of
their efficacy. For example, in a comparative study on
polytetrafluoroethylene (PTFE) tubes, contaminated by a liquid
medium inoculated with Pseudomonas aeruginosa, using five different
alternative disinfectant solutions: two peracetic acid solutions
(with and without an activator), glutaraldehyde, orthophthaldehyde
and succine dialdehyde, it was shown that repeated treatments of a
PTFE tube with a 2% glutaraldehyde solution induced an important
accumulation and/or fixation of protein, compared to
peracetic-acid-based disinfectants, for which the accumulation
and/or fixation of protein remained low, but varied from one
formulation to another (L. Pineau, C. Desbuquois, B. Marchetti, D.
Luu Duck, "Comparison of the fixative properties of five
disinfectant solutions", The Journal of Hospital Infection, Vol.
68, Issue 2, Pages 171-177 (February 2008).
In this context again, the enzyme Trehalase can be included in
various formulations of disinfection solutions, and can be used as
directed in the previous embodiments for composition and methods
for treatment of biofilm-based infections on medical devices.
It should be understood that the copper may operate to short
circuit the current in the cell membrane to disturb the "trans
membrane potential" and can cause oxidative damage and thus,
"punch" a hole in the bacterium. It is possible to add Trehalase to
Medicinal Copper, and even possible to apply Trehalase and copper
in a spray as an adhesive and spray it onto surfaces. Besides
copper, it is also possible to use Trehalase in combination with
biofilm disruptors such as xylitol, lactoferrin, and D-amino acids.
Xylitol is a sugar alcohol used as a sweetener.
The bacterial biofilms are the major contributing factors to
chronic wounds, such as diabetic foot ulcers, pressure ulcers, and
venous leg ulcers, providing increased bioburden to the wounds and
interfering with the healing process. Traditional methods of
treatment have proven ineffective, therefore therapy of non-healing
wounds demands biofilm-targeted strategies.
Studies using a colony-drip-flow reactor biofilm model,
demonstrated positive result of the combined treatment of bacterial
biofilms with the innate immune molecule lactoferrin and the rare
sugar-alcohol xylitol against a clinical wound isolate; for both a
single species biofilm and a dual species biofilm, the
lactoferrin/xylitol hydrogel in combination with the silver wound
dressing Acticoat.TM. had a statistically significant reduction in
biofilm cell viability compared to the commercially available wound
hydrogel (Ammons M C, Ward L S, James G A, "Anti-biofilm efficacy
of a lactoferrin/xylitol wound hydrogel used in combination with
silver wound dressings", Int Wound J, 2011 Jun. 8(3); 268-73). In
an in vitro biofilm model with a clinical isolate of P. aeruginosa,
subjected to treatment with either lactoferrin or xylitol alone or
in combination, it was shown that combined lactoferrin and xylitol
treatment disrupted the structure of the P. aeruginosa biofilm and
resulted in a >2 log reduction in biofilm cell viability; in
situ analysis indicated that xylitol appeared to disrupt the
biofilm structure and lactoferrin increased the permeability of
bacterial cells (Ammons M C, Ward L S, Fisher S T, Wolcott R D,
James G A, "In vitro susceptibility of established biofilms
composed of a clinical wound isolate of Pseudomonas aeruginosa
treated with lactoferrin and xylitol", Int J Antimicrob Agents,
2009 March, 33(3):230-6.).
As noted before, the enzyme trehalase can be used in combination
with antimicrobials (ex, silver gel or silver-based wound
dressings) for chronic wounds treatment along with other
antimicrobials, or can be used in combination with lactoferrin or
lactoferrin/xylitol to increase effectiveness of these treatment
modalities. As further noted before, to increase the healing
process of such chronic wounds, trehalase in combination with
antimicrobials can be used along with antioxidants,
anti-inflammatory and immune-modulating substances. Trehalase may
be added in the preparation and amounts disclosed previously.
In many cases, chronic wounds have tendency to be additionally
contaminated by Gram-positive pathogens, such as S. aureus and
Streptococci spp. that contribute to increased bioburden in mixed
species biofilms. In these cases, Trehalase in combination with
polymicrobial antibiotics and/or natural antimicrobials (i.e.,
silver compounds as noted before) can provide dual effect,
targeting the biofilm structure and reducing viability of
biofilm-forming pathogens (including both biofilm-embedded cells
and dispersed planktonic cells).
It is also possible to add units of Trehalase enzyme to an
antibiotic ointment that may include one or more of bacitracin
zinc, neomycin sulfate that may be equivalent to a neomycin base
and a polymyxin B sulfate. Although the amounts can vary in one
example, 300 to 500 units of a bacitracin zinc may be added to 2.5
to 4.5 mg of the neomycin base and 4,000 to 6,000 polymyxin B units
added as the polymyxin B sulfate together with units of Trehalase
with sufficient units of activity and in one example, 0.2 to 1.0
units per mg and around 0.4 units per milligram.
A sufficient amount of Trehalase may be added to an enzyme
supplement that could include protease as either 4.5, 6.0 and/or
3.0 such as respective amounts of 14,000 to 23,000 HUT for the
protease 4.5, 2,000 to 4,000 HUT for the protease 6.0, and 13 to 19
SAPU for protease 3.0 and in specific amounts about 19,000; 3,000;
and 16. About 3,000 to 5,000 DU of amylase may be added and in a
specific amount 4,200, and 7 to 13 AGU of glucoamylase and in a
specific amount 10 AGU, 20 to 28 LU of lipase and in a specific
amount 24 LU. 90 to 150 CU of cellulase may be added (specific 120
CU) and 180 to 260 SU of invertase (specific amount 220 SU), and
500 to 600 DP for malt diastase (specific 550 DP). There may also
be included about 80 to 120 GALU (100 GALU) for alpha-galactosidase
and 1,600 to 2,400 HUT of peptidase (specific amount 2,000 HUT).
Many of these quantities can vary and be supplemented with other
enzymes depending on the end use desired. The Trehalase can be
supplemented in an effective amount of 0.2 to 1.0 unit in 1 mg of
protein. It may also be added with probiotics such as a blend of
Lactobacillus acidophilus and biphobacterium lactis such as around
4 billion for the probiotic in one non-limiting example.
It is also possible to combine the Trehalase with antimicrobial
peptides that may be an alternative to small molecule antibiotics,
including modifications of short antimicrobial peptides. This could
include lipopeptides and a combination of fatty acids with cationic
antimicrobial peptides that help change the confirmation of the
antimicrobial peptides secondary structure in the presence of
bacterial membranes.
It is possible assay Trehalase based on the technique described in
Dahlquist, A., (Assay of Intestinal Disaccharidases Analytical
Biochemistry (1968)). An enzymatic assay of Trehalase has been
formulated as (EC 3.2.1.28) by Sigma-Andrich.
Principle: Trehalose+H.sub.2O.sup.Trehalase>2 Glucose
Conditions: T=37.degree. C., pH=5.7, A.sub.340 nm, Light path=1
cm
Method: Spectrophotometric Stop Rate Determination
The Reagents in this assay include:
A. 135 mM Citric Acid Buffer, pH 5.7 at 37.degree. C., which is
prepared 100 ml in deionized water using Citric Acid, Free Acid,
Monohydrate, as Sigma Prod. No. C-7129, corresponding to the citric
acid monohydrate. It is adjusted to pH 5.7 at 37 C with 1 M
NaOH.
B. 140 mM D-Trehalose Solution, which is prepared by 10 ml in
Reagent A using D(+)Trehalose, Dihydrate, as Sigma Prod. No. T-5251
corresponding to D-(+)-Trehalose dehydrate
C. 500 mM Tris Buffer, pH 7.5 at 37.degree. C., which is prepared
by 100 ml in deionized water using Trizma Base, as Sigma Prod. No.
T-1503. It is adjusted to pH 7.5 at 37 C with 1 M HCl.
D. Trehalase Enzyme Solution. Immediately before use, prepare a
solution containing 0.1-0.3 unit/ml of Trehalase in cold Reagent
A.
E. Glucose Determination Vial. It is possible to use an ion
exchange type as Sigma Stock No. 16-10 (Mono QHR 16/10), Glucose
(HK) 10 Reagent. The contents are dissolved in 10 ml of deionized
water.
Procedure:
Step 1: Pipette (in milliliters) the following reagents into
suitable cuvettes:
TABLE-US-00003 Test Blank Reagent A (Citrate Buffer) 0.3 0.3
Reagent D (Enzyme Solution) 0.1 0.1
Mix by inversion and equilibrate to 37.degree. C. using a suitably
thermostatted spectrophotometer. Then add:
TABLE-US-00004 Reagent B (D-Trehalose) 0.1 --
Immediately mix by inversion and incubate at 37.degree. C. for
exactly 15 minutes. Then add:
TABLE-US-00005 Reagent C (Tris Buffer) 0.5 0.5 Reagent B
(D-Trehalose) -- 0.1
Step 2: Pipette (in milliliters) the following reagents into
suitable cuvettes:
TABLE-US-00006 Test Blank Reagent E (16-10) 3.0 3.0
Equilibrate to 37.degree. C. Monitor the A.sub.340 nm until
constant, using a suitably thermostatted spectrophotometer. Record
the initial A.sub.340 nm for both Test and Blank. Then add:
TABLE-US-00007 Test Solution 0.1 -- Blank Solution -- 0.1
Immediately mix by inversion and record the increase in A.sub.340
nm until complete (approximately 5 minutes). Obtain the final
A.sub.340 nm for both the Test and Blank. Calculations:
.DELTA.A.sub.340 nm Test=A.sub.340 nm Test Final-A340 nm Test
Initial .DELTA.A.sub.340 nm Blank=A.sub.340 nm Blank
Final-A.sub.340 nm Blank Initial Units/ml enzyme=(.DELTA.A.sub.340
nm Test-A.sub.340 nm Blank)(1.0)(3.1)/(6.22)(2)(15)(0.1)(0.1)
6.22=Millimolar extinction coefficient of .beta.-NADH at 340 nm
2=Number of Glucose molecules per molecule of Trehalose 15=Reaction
time (in minutes) of Step 1 1.0=Final volume (in milliliters) of
Step 1 3.1=Final volume (in milliliters) of Step 2 0.1=Volume From
Step 1 used in Step 2 0.1=Volume (in milliliters) of enzyme used
Units/mg protein=units/ml enzyme/mg protein/ml enzyme UNIT
DEFINITION: One unit will convert 1.0 .mu.mole of trehalose to 2.0
moles of glucose per minute at pH 5.7 at 37.degree. C. (liberated
glucose determined at pH 7.5). FINAL ASSAY CONCENTRATION: In a 0.50
ml reaction mix, the final concentrations are 135 mM citric acid,
28 mM D-trehalose, and 0.01-0.03 unit of trehalase.
The previous description of the disclosed embodiments is provided
to enable any person skilled in the art to make or use the
disclosed embodiments. Various modifications to these embodiments
will be readily apparent to those skilled in the art, and the
principles defined herein may be applied to other embodiments
without departing from the scope of the disclosure. Thus, the
present disclosure is not intended to be limited to the embodiments
shown herein but is to be accorded the widest scope possible
consistent with the principles and novel features as defined by the
following claims.
* * * * *
References